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Chapter 1: Mechanisms for DNA Charge Transport · DNA. Poor base stacking, associated with this...
Transcript of Chapter 1: Mechanisms for DNA Charge Transport · DNA. Poor base stacking, associated with this...
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Chapter 1:
Mechanisms for DNA Charge Transport†
† adapted from Genereux, J.C.; Barton, J.K. Chem. Rev. 2009, in press.
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1.1. INTRODUCTION
DNA charge transport (CT) chemistry has received considerable attention by
scientific researchers over the past 15 years since the first provocative publication on
long-range CT in a DNA assembly.1,2 This interest, shared by physicists, chemists and
biologists, reflects the potential of DNA CT to provide a sensitive route for signaling,
whether in the construction of nanoscale biosensors or as an enzymatic tool to detect
damage in the genome. Research into DNA CT chemistry began as a quest to determine
whether the DNA double helix, a macromolecular assembly in solution with -stacked
base pairs, might share conductive characteristics with -stacked solids. Physicists
carried out sophisticated experiments to measure the conductivity of DNA samples, but
the means to connect discrete DNA assemblies into the devices to gauge conductivity
varied, as did the conditions under which conductivities were determined. Chemists
constructed DNA assemblies to measure hole and electron transport in solution using a
variety of hole and electron donors. Here, too, DNA CT was seen to depend upon the
connections, or coupling, between donors and the DNA base-pair stack. Importantly,
these experiments have resolved the debate over whether DNA CT is possible. Moreover
these studies have shown that DNA CT, irrespective of the oxidant or reductant used to
initiate the chemistry, can occur over long molecular distances but can be exquisitely
sensitive to perturbations in the base-pair stack.
Here we review some of the critical characteristics of DNA charge transport
chemistry, taking examples from a range of systems, and consider these characteristics in
the context of their mechanistic implications. This chapter is not intended to be
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exhaustive but instead to be illustrative. For instance, we describe studies involving
measurements in solution using pendant photooxidants to inject holes, conductivity
studies with covalently modified assemblies, and electrochemical studies on DNA-
modified electrodes. We do not focus in detail on the differences amongst these
constructs but instead on their similarities. It is the similarity among these various
systems that allows us to consider different mechanisms to describe DNA CT. Thus we
review also the various mechanisms for DNA CT that have been put forth and attempt to
reconcile these mechanistic proposals with the many disparate measurements of DNA
CT. Certainly the debate among researchers has shifted from “is DNA CT possible?” to
“how does it work?” This chapter explores this latter question in detail.
1.2. PROPERTIES OF LONG-RANGE CHARGE TRANSPORT IN DNA
Among the most interesting characteristics of charge transport in DNA is the long
distance over which it occurs (Figure 1.1).3-6 Nevertheless, there are some DNA systems
that do not mediate charge over long distances. How DNA CT occurs depends upon
coupling and the structure and dynamics of the DNA assembly. The chemistry and
photophysics of the photoexcited acridine (Acr*+) containing systems, which mediate CT
over only a few base pairs, have been particularly well-characterized in this regard.7,8 It is
important to note that the same physical laws apply to all CT processes.9 The essential
distinctions are with respect to the relative roles which different mechanisms play, and it
is in this respect that long-range CT, with effective coupling to the base-pair stack, differs
from short-range CT, with poor coupling. Here we focus on long-range CT, where
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Figure 1.1. Transverse and longitudinal perspectives of DNA. The sugar phosphate
backbone envelops the hydrophobic base pairs. The planar base pairs form a one-
dimensional -stack down the center of the DNA, insulated by the backbone.
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transport is through the base pair stack, and discuss short-range systems only to the
necessary extent to clarify the distinctions between the two regimes.
1.2.1. COUPLING TO THE DNA
It is notable that initial measurements of DNA-mediated charge transport for both
photooxidation experiments10 and device experiments11 found rates and conductivities
spanning several orders of magnitude over comparable distances, depending on the
experimental conditions. This foreshadowed the same observation in scanning tunneling
microscopy (STM) measurements of conductivity through other molecular bridges,12 and,
ultimately, was for the same reason. For short molecular bridges, it has been established
both experimentally12 and theoretically13 that the coupling between the bridge and the
donor (or acceptor) can dominate the observed conductivity. Similarly, when DNA is the
bridge, the coupling can have a dramatic effect on both charge transport rates and yields
(Figure 1.2).14-18 Characteristically, conductivity measurements that have not provided
covalent contact between the DNA and the device yield a spectrum of behavior: from
insulating to superconductive.11,19-23
In the case of DNA, the essential coupling is into the -stack of the bases. This is
a marked challenge, as DNA is essentially “insulated”, with sugars and phosphates
flanking the periphery of the bases.24 This insulation, in part, explains why early
experiments on dry DNA found insulating behavior, in contrast to that observed with
conducting organic polymers. A series of well-conjugated charge donors and acceptors
are now employed by various groups,25,26 including metallointercalators, organic
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Figure 1.2. DNA-mediated CT requires electronic coupling to the base pair stack.
(A) Electrochemical reduction of an electronically well-coupled anthraquinone (AQ) is
facile, while that of a poorly coupled AQ is suppressed.18 (B) MutY competently reduces
an oxidized nitroxide spin label that is well coupled to thymidine, but not the nitroxide
conjugated through the partially unsaturated linker.160 (C) For a series of polypyridyl
RuIII(bpy)2L ground state oxidants, the yield of oxidative damage to DNA scales with the
size and planarity of the intercalating ligand.15
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intercalators, organic end-cappers, and modified bases. In several cases, direct
comparison has been made between similar photooxidant pairs that differ primarily in
their ability to couple well with the base stack. These examples include the adenine
analogues ethenoadenine and 2-aminopurine (Ap),14 two different coupling strategies for
ethidium bromide,27 and, most notably, a series of intercalating ruthenium analogues with
decreasing planarity in the intercalating ligand.15 As an extreme case, for two ruthenium
complexes that are unable to intercalate, and that are attached on opposite ends of a short
DNA duplex via terminal phosphate modification, the CT rate was found to be ~10-6 s-1;28
this is what would be expected for the rate were the metal complexes connected solely
through their tethers. Similarly, electrochemiluminescence studies find the same rates
for DNA-mediated CT between a DNA-modified gold electrode and tethered Ru(bpy)32+
as are observed through solely the tether itself.29 In electrochemical studies of methylene
blue covalently attached to a DNA duplex, effective transport is found only when the
methylene blue is stacked in the helix, not under high salt conditions, where the dye,
although still linked by a -bonded tether, is unstacked.30 Indeed, electrochemical
measurements on DNA films generally have been shown to be rate-limited by tether
linking the DNA to the electrode surface31. In each case, it is clear that the coupling
between the donor/acceptor pair and the bridge is dominating the measurement, and that
the bridge is the -stack of DNA.
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1.2.2 GLOBAL STRUCTURAL INTEGRITY
The structure of DNA is central to its extraordinary effectiveness as the genetic
template for the cell. This relationship between structure and function is underscored by
the extent of the biological function that was first predicted in the landmark papers that
reported the proper three-dimensional structure.32,33 Hence, it is not surprising that DNA-
mediated CT is also substantially affected by the global structure of a DNA sample. This
is clear when considering the results of conductivity measurements on single or few DNA
strands that have been performed in recent years. Various measurements from 1996 to the
present have found DNA conductivities covering several orders of magnitude.
Furthermore, conductivity has been found to be dependent on sequence, hydration,
length, temperature, and hybridization in some experiments, while independent of each of
those in others. Ultimately, the vast differences in observations can be largely reconciled
by comparing the sample preparation methodologies of the individual studies.11
Conditions that cause global DNA conformational changes or damage can both increase
or decrease the observed conductivity. In one extreme case, it was found that imaging
conditions commonly used prior to conductance measurements lead to a morphological
change in the structure of the DNA, that is itself correlated with increased conductivity.20
Among experiments that examine undamaged DNA, a profound difference is
always observed between single-stranded and double-stranded DNA: single-stranded
DNA does not mediate CT over long distances. This has been observed by direct
conductivity studies,34 photooxidation,35 transient absorption,36 electrochemical AFM,37,38
STM,39 electrogenerated chemiluminescence,29 and electrochemical experiments in DNA
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monolayers.40 The caveat in interpreting studies on single-stranded DNA is, however,
that its structure and, importantly, stacking are heterogeneous and extremely dependent
on sequence.
DNA, stabilized by a variety of hydrophobic and hydrophilic interactions and
evolved for an aqueous environment, undergoes gross structural changes as a result of
moving from a hydrated to a dehydrated environment.41 Critically, these changes are to
the equilibrium conformation of DNA; the effects of dehydration on DNA dynamics are
not well understood. Highly bound waters play a major role in the dynamics that gate
molecular recognition and other biochemical interactions between macromolecules.42
Regrettably, the first and many recent measurements of DNA conductivity were
performed under vacuum. Vacuum is ideal for conductivity measurements due to the
suppression of voltage leak and the associated background current. Even experiments
performed in the presence of water frequently deposit the DNA under vacuum conditions.
Similar to the previous case of poorly coupled versus well-coupled systems, there is wide
disparity between the conductivities observed under conditions of low humidity.
Recently, progress has been made in understanding the role of humidity in many of the
poorly coupled systems.43 Even without strong coupling into the DNA base stack, water
adsorbed on the DNA and in DNA bundles can mediate ionic conduction. The amount of
adsorbed water will depend strongly on humidity, and also on the adsorption environment
of the DNA. This helps explain why many systems in which coupling to DNA was poor
were still observed to conduct.19,23,44
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Not surprisingly, experiments that have preserved the DNA in its native
conformation, with leads covalently coupled to the bridge, have shown remarkably
similar (and substantial) conductivities (Figure 1.3).34,37,44,45 The conductivity measured
by Xu et al. across a dodecanucleotide with terminal propylthiol-Au contacts (> 40 Å) is
comparable (6 x 10-4 G0, where G0 is the quantum unit of conductance) to that found
across the much smaller benzenedimethanethiol (~10 Å) under the same experimental
conditions,47 though this comparison is complicated by the possibility that DNA
accommodates internal stretching during the measurement rather than extruding gold
from the molecular junction, as is postulated for benzenedimethanethiol.
As is the case with water, ionic strength can dictate the conformation of DNA.
High ionic strength drives the transition from B-form to the more extended Z-form of
DNA. Poor base stacking, associated with this condensed structure, leads to less efficient
DNA-mediated CT.48 Conversely, sufficiently low ionic strength leads to strand
dehybridization, which also suppresses CT.
Beyond issues of ionic strength, there is conflicting evidence as to whether the
identity of the counterion affects CT. Some calculations have shown that counterion
identity does not affect the electric field inside the DNA,49 while others have found that
movement of a single sodium has profound effects on base energies.50 Similarly
contrasting results have been observed in experimental work.51-54 For solvent-exposed
donors and acceptors, an ion-pair can form between dye and counter-ion that itself
profoundly affects CT rate and yield.55
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Figure 1.3. Devices for measurement of single molecule DNA conductivity through
molecular contacts. In each case, currents between 10 and 100 nA are obtained for
modest source-drain and gating voltages. A) A gold nanoparticle allows strong coupling
between the EC-AFM tip and an individual 26mer DNA molecule on a gold electrode.37
B) The gold STM tip is slowly brought in contact with thiol-modified DNA (8mer),
allowing a histogram of conductance over many different orientations.45 C) A single
15mer DNA is covalently attached across a gap between single-walled carbon
nanotubes.34
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More chemically controlled experiments elucidate the structural basis of
environmental effects. RNA/DNA hybrids and double stranded RNAs adopt the A-form
while alternating purine-pyrimidine sequences under certain conditions adopt Z-form
structures. Both conformations support DNA charge transport, though Z-form is an
inferior bridge relative to A-form and B-form for electrochemical,48 but not
photooxidation assays.56 Not surprisingly, the competence for mediating CT has been
shown to follow the extent of base stacking, both in solution studies with Ap as the
photooxidant57 and in electrochemical experiments monitoring the efficiency of reducing
an intercalated redox probe.48 Again, different coupling of the redox probes into these
different conformations means that they cannot be quantitatively compared.
Perhaps most interesting is the comparison of rates of intrastrand versus
interstrand base-base CT in DNA assemblies modified with Ap.14,35,57 Here for the B-
conformation, intrastrand CT is found to be three orders of magnitude faster than
interstrand CT, consistent with the fact that stacking in the B-conformation is exclusively
intrastrand; CT across strands requires CT across a hydrogen bond. However, in the A-
form there is a mix of interstrand and intrastrand stacking down the helix, and here we
observe that rates of intrastrand and interstrand CT are comparable.
1.2.3. LOCAL STRUCTURAL INTEGRITY
A variety of studies have found similar effects of disrupting the base stack locally.
The assays include electrochemical experiments in both films30,58,59 and devices,34 and
solution experiments using time-resolved fluorescence,60 irreversible trapping of
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chemical product61 and transient absorption measurements.62 The presence of mismatches
lowers both the rate and yield of DNA-mediated CT, and the extent of this attenuation
scales with the base pair lifetime.63 Abasic sites64 and destabilizing lesions65 also interfere
with CT through DNA films.
Regarding those experiments that utilize product trapping, however, it is
important to note that the results are convoluted with two effective clocks. The first is the
rate of back electron transfer (BET), if it occurs.66 The second is the rate of product
trapping.66-69 A disruption of the -stack will only be observable in product trapping
experiments if it is sufficient to disrupt equilibration of the radical cation on the time
scale of BET and product trapping.70 Towards this end, guanine damage assays have
recently been replaced by assays for fast decomposition of a radical trap. N-
cyclopropylguanosine (CPG),71,72 N-cyclopropyladenosine (CPA),17,73,74 and N-
cyclopropylcytidine (CPC)75,76 are synthetically accessible, cause minimal perturbation to
the DNA-duplex, and undergo irreversible picosecond ring-opening upon oxidation or
reduction.
Not all modifications of DNA suppress long-range CT. A dephosphorylation nick
to the backbone, despite causing substantial change to local ion density, does not have a
measurable effect.77,78 Furthermore, some modifications can enhance CT. Most notably,
DNA with adenines replaced by the lower-potential base deazaadenine or the better-
stacking benzodeazaadenine improves the rate and yield of DNA-mediated CT.79-81
In addition to global changes in structural integrity, subtle modulations to
structure can also have profound effects on the rates and yields of DNA-mediated CT.
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DNA-mediated CT is attenuated by the presence of mismatches, even though mismatched
base pairs cause only minor distortions to the structure of DNA.82,83 Nevertheless,
mismatch discrimination has been observed in charge transport through DNA films,58,59,84
single molecule devices,34,85 and photooxidation systems.35,61 This attenuation is identical
for oxidative and reductive CT,86 implying mechanistic similarity. Importantly, the extent
of mismatch discrimination corresponds to the base-pair lifetime associated with the
specific mismatch.63 In the extreme case, an abasic site completely suppresses CT.64,87,88
Subtle lesions, such as the oxidative guanine products O6-methyl-guanine and
8-oxoguanine, attenuate CT.65 It should be noted that although 8-oxoguanine terminates
DNA-mediated CT as a thermodynamic and kinetic trap at high driving force,89,90 this
and other damaged base products also attenuate CT even under driving forces
incompetent for direct oxidation. This property of DNA-mediated CT has led to the
development of a new class of DNA-detection devices,91,92 and might be relevant to
damage detection in the cell.93
Local changes to structure can also be induced. Inside the cell, proteins can bend,
twist, and dehybridize DNA, and some can extrude bases as well. Not surprisingly, many
of these binding events have severe effects on DNA-mediated CT. Monitoring DNA-
mediated electrochemistry to SoxR, a transcription factor that initiates the oxidative stress
response in E. coli, is consistent with the prediction that the oxidized form twists DNA to
initiate transcription,94 as has since been validated by a recent crystal structure.95 CT is
also inhibited by the sequence-specific binding of TATA-binding protein (TBP),58,96
which bends DNA. One particularly informative experiment involved the methylase
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M.HhaI, which extrudes a cytidine within its recognition sequence, replacing it with
glutamine. DNA with the HhaI recognition site shows attenuated CT in the presence of
the protein. The Q237W mutant, in which the intercalating glutamine is replaced with the
aromatic ligand tryptophan, however, barely affects CT compared to the absence of
protein.58,97 Importantly, proteins that do not distort the DNA -stack, such as
antennapedia homeodomain protein or unactivated R.PvuII, do not attenuate DNA-
mediated CT.97 Indeed, the rigidification associated with R.PvuII binding increased CT
through the dynamically flexible TATA binding site. An interesting exception is the case
of R.BamHI, a restriction endonuclease which does not bend the DNA -stack, but
contains an asparagine guanidinium that hydrogen bonds to a guanine in its cognate site.
It has been shown that R.BamHI attenuates DNA CT to guanines both within and beyond
its binding site, presumably due to electrostatic modulation of the intervening DNA via
the hydrogen bonding interactions.98
Protein binding can also affect the fate of radicals in DNA.99-101 Guanine radical
has been shown to crosslink to histones and short peptides. In one case,102 excitation of
an anthraquinone (AQ)-DNA conjugate bound to a reconstituted nucleosome particle led
to DNA-protein crosslinks; experiments with a different photooxidant with facile back
electron transfer found no such effect.103 Since it is clear that migration can occur over
long distances in both isolated nuclei104 and mitochondria,105 this might not be a fast
pathway for radical quenching; the time scale of protein-DNA crosslinking in the
presence of guanine radical is similar to that for guanine radical decomposition to 8-
oxoguanine.106 These crosslinks are reversible under the processing conditions used to
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convert base damage to strand cleavage, and hence are likely hidden in typical gel
analysis experiments. Furthermore, protein cross-links can be formed as a secondary
product after radical degradation in DNA to 8-oxoguanine.107 Recently, it has been shown
that a protein that disrupts the -stack, Hbb of Borrelia burgdorferi, affects the
conversion of radical damage to interstrand crosslinks.108
DNA CT is sensitive to even more subtle deviations in stacking integrity. The
strongest stacking interactions occur between consecutive purines, as has been shown
both experimentally and computationally. Extended purine-pyrimidine runs correspond to
the minimal extent of base-stacking, while purine-purine runs, particularly adenine tracts,
correspond to the maximal extent for DNA containing natural nucleotides. This
relationship is borne out in the sequence dependence of CT.51,109-111 Note that in
photoactivated studies of electron transport, runs of pyrimidines, which are more easily
reduced, are the preferred sequences.112 Likely here too, the decreased flexibility of
homopurine-homopyrimidine sequences plays a role.
1.2.4. CONFORMATIONAL GATING
The rate and efficiency of charge transfer is centrally related to the structure of the
individual pathway(s) that mediates CT between the donor and acceptor. Over long
distances, however, it is inevitable that fluctuations will be induced at nonzero
temperature, such that the equilibrium structure only reflects an average over the
ensemble. If these fluctuations are sufficiently large, and slower than CT for the
equilibrium conformation, then CT no longer is properly described by a unitary rate
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constant, but will instead proceed with a complicated time-dependence that convolutes
the conformational dynamics with the CT rates of the different conformations.113 To
avoid this, the best-performing molecular wires are chosen for, among other properties,
rigid homogeneous conformations.114 For example, the conduction of certain
oligophenyleneethynylenes can be substantially enhanced by the presence of bulky side-
groups that limit the conjugation-breaking rotation around the acetylene bonds.115,116
It is not surprising that DNA, which even as a relatively short 15-mer contains
several hundred atoms, exhibits substantial conformational flexibility. Interestingly, the
effect of conformational gating in DNA is generally to increase CT rather than to
decrease it. Duplexes frozen in glass show no attenuation in CT between the
photooxidant Ap and neighboring guanine hole acceptor (Figure 1.4).117 The insertion of
a single adenine, however, between donor and acceptor completely suppresses CT. The
temperature-dependence of CT between Ap* and G for varying bridge length has been
studied by both femtosecond transient absorption spectroscopy35 and steady-state
fluorescence quenching measurements.54 The temperature-dependence has also been
measured for CT between tethered, photoexcited [Rh(phi)2(bpy')]3+ and CPG.68 For all
bridges, CT efficiency increases with temperature, and the temperature dependence is
greater with increasing bridge length.
Although thermal activation on its own does not imply a mechanism, the
sequence dependence establishes a strong relationship between bridge structure and the
activated process. The temperature dependence of the electronic contribution to the rate
should be the same irrespective of increasing bridge length. The temperature dependence
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Figure 1.4. The sequence and temperature dependence of single-step oxidation of
guanine by photoexcited 2-aminopurine (Ap) in DNA.35,54,117 CT yield in well-matched
Ap(A)4G increases with temperature (+ ), up to duplex melting. Two perturbations that
disturb CT due to poor stacking dynamics, an A-A mismatch and the sequence ATAT,
attenuate CT at room temperature but are comparable to the A4 sequence at higher
temperature, while CT through a perturbation that disturbs CT due to an electronic
barrier, AAIA, is only partially recovered at high temperature. This argues that the CT
activation is related to the flexibility of the bridge. At low temperature (77K), an
intervening adenine eliminates CT from Ap to G, implying that the equilibrium
conformation is not the CT active conformation.
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of the nuclear contribution, due to bridge-length dependences on driving force and
reorganization energy (see below), can naturally be bridge-length dependent, but should
disappear for distances above 10 to 15 Å,118 which are exceeded here. Conformational
gating to reach a CT-active state, however, is expected to lead to increased CT with
temperature, and for this increase to be greater when more bridge units are required to
align, i.e., for longer bridges.
Interestingly, this increase in the rate with respect to increasing temperature is
even more profound when the bridge is poorly stacked ATAT,54 or contains an AA
mismatch,35 consistent with the model whereby these sequences disrupt DNA due to poor
dynamic stacking (Figure 1.4). When the bridge is AAIA, which stacks well but
attenuates CT from Ap* due to the inosine potential barrier, the increase in CT with
respect to temperature is only modest. These experiments strongly suggest that the
equilibrium conformation of DNA is not the active conformation for long-distance CT,
but that conformational gating allows the formation of CT-active states. This is in direct
contrast to the usual role dynamic disorder plays in molecular wires, where distortion
from the equilibrium conformation decreases coupling and transport.119
There are two different implications of conformational gating. In one sense,
reorientation of the photooxidant with respect to the DNA to form a CT-active
conformation can be rate-limiting. Alternatively, formation of a CT-active state in the
DNA itself can be the rate-limiting step for CT. Both senses are represented by the case
of ethidium bromide. Ethidium bromide (Et+) is competent for DNA-mediated oxidation
and reduction of deazaguanine (ZG) and [Rh(phi)2(bpy')]3+ respectively.27,60,120,121
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Femtosecond transient absorption and fluorescence up-conversion spectroscopy of Et+
site-selectively intercalated in DNA found that the Et+ oxidized ZG with two rate
constants.120 One (5 ps) corresponds to Et+ that is already present in a CT-active
conformation. The other (75 ps) corresponds to the gating of Et+ to reach a CT-active
conformation with respect to the DNA. This is the first sense of conformational gating.
As the distance between Et+ and ZG increases, the rate of each component is unaffected,
but the amplitudes monotonically decrease, suggesting that the increase in distance
lowers the yields by virtue of changing the population of CT-active states, rather than by
affecting the inherent rates of CT through the DNA. This represents the second sense of
conformational gating.
To further test this model, a new method of conjugating Et+ to DNA as a rigid
base-pair surrogate was developed (Figure 1.5).27 For the Et+ separated from the hole
acceptor, ZG, by a single base pair, the rate is similar to that for CT from the intercalated
Et+. This rate drops four orders of magnitude if another adenine is inserted between the
Et+ and ZG. Beyond a distance of two base pairs, the rate is constant. The authors
interpreted this system as one that held the Et+ rigidly in a CT-inactive state. The rate-
limiting step is injection, which for a poorly coupled donor exhibits a steep distance
dependence. At sufficient donor-acceptor separation, re-orientation of the donor is
competitive with the slow CT from poorly coupled Et+. Now the apparent distance-
dependence is flat, for the same reason as for the intercalating Et+.
A similar explanation might serve for the slow rate of charge injection into
stilbene-capped hairpins where several AT base pairs separate the photoexcited stilbene
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Figure 1.5. (A) The rate of coherent deazaguanine (ZG) oxidation by ethidium bromide is
the same over short distances for both the flexible linkage (Et1, black, both CT rates
shown) and the rigid linkage (Et2, gray).27 For two intervening nucleotides, a sharp drop
in rate is observed for the rigid Et+, but the rate is unaffected for the flexible Et+. (B) This
steep drop in rate over short distances is consistent with that observed for oxidation of
guanine by a photoinduced sugar radical (circles),207 and CT between hairpin capping
stilbenes (squares, photooxidation of Sd by Sa),125 and has been attributed to a crossover
between coherent superexchange and incoherent hopping. In the latter case, comparison
of injection and hole arrival rates supports superexchange for one or two intervening base
pairs, and hopping for three or more base pairs between the stilbenes. Adapted from
References 27, 207, and 125.
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hole donor from a guanine hole acceptor. Stilbene-4,4'-dicarboxamide incorporated as a
bridging and capping element in short AT containing hairpins (Sa-AT) has an extended
nanosecond lifetime and hence higher fluorescence quantum yield versus the free dye,
but a single GC pair close to the stilbene dramatically decreases the fluorescence
intensity, via CT quenching.122,123 The robust fluorescence in the absence of guanine was
taken as evidence for a lack of CT between stilbene and adenine, despite the presence of
several low-yield picosecond decay components. Eventually, these components were
assigned to CT between the excited stilbene and adenine.124,125 The charge-separated state
can either undergo recombination, or, in the presence of distal guanine or a lower energy
stilbene-4,4'-diether (Sd), the hole migrates to the acceptor leading to CT quenching.126 It
was argued that the recombination recovers the excited state, and that hence charge
injection in the Sa-AT constructs only minimally quenches fluorescence.124 This is
distinct from exciplex emission, as the emission spectrum is similar to the unconjugated
fluorophore. The stilbene radical anion is solvent exposed, and the motions of DNA,
associated counterions, and bound water allow rapid relaxation of dyes,127 so it is
unlikely that recombination-induced emission would be of the same energy as radiation
from the initial excited state. An alternate explanation is that this fast injection is limited
to a small population that is in a CT-active conformation. The remaining population
fluoresces normally in the absence of nearby guanine. In that context, the slow, strongly
distance-dependent direct charge transfer from excited stilbene to guanine can be
interpreted in the context of a donor that is not in a CT-active state with respect to the -
stack.122
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1.2.5. BACK ELECTRON TRANSFER
Inevitably, photo-induced charge separation events are followed by charge
recombination, also termed back electron transfer (BET). After all, if charge
recombination is thermodynamically unfavorable, then charge separation is
thermodynamically favorable and will not require photoexcitation. The effect of BET
varies by the nature of the assay. Assays for the presence of the charge separated state,
such as the slow oxidation of guanine cation radical, will generate yields that are
convoluted with BET processes. In two extreme cases, thionine128 and Ap,73 which are
competent for efficient charge separation, are not competent for the formation of
permanent guanine oxidation products.
The case of the two excited electronic states of AQ offers a nice comparison of
photooxidation with fast and slow BET. Irradiation of DNA-conjugated AQ at 350 nm
promotes it to the singlet excited state, which relaxes to the triplet state. Both states are
competent for direct oxidation of all four bases in DNA, but only the triplet radical anion
reduces oxygen to superoxide. The singlet radical anion undergoes rapid BET,
regenerating the initial state, while the charge injected by the triplet radical anion is
persistent, and can equilibrate along the DNA on a longer time scale.129,130 This scheme
explains the incompetence of AQ to oxidatively repair cyclobutane thymine dimer;131,132
repair can only proceed from the singlet state.
Experiments that rely on slow product trapping at guanine need BET to provide
the fundamental clock that allows discrimination of CT attenuation.66 Hence, although
the results will be qualitatively diagnostic, the quantitative accuracy will only hold
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relative to the BET rate for that system. For example, a single negatively charged
phosphate group near the intercalation site of a tethered rhodium photooxidant changes
the observed ratio of damage between a distal and proximal GG site by an order of
magnitude, indicating that the distal/proximal damage ratio is not solely determined by
the intervening bridge.52
Short-range CT is particularly subject to BET, as the recombination has a steeper
distance dependence than separation, most likely due to greater separation in donor-
bridge-acceptor energies, as discussed in Section 1.3.2.122 This was first exploited in
guanine damage systems using AQ as the donor, but has since been systematically
studied in Acr+-phenothiazine (Ptz) and napthalimide (NI)-Ptz systems.133-136 In the
former, suppressing BET allowed the extension of a canonical short-range CT system
into one that exhibited persistent CT separation over a long range!136
1.2.6 INJECTION AND MIGRATION EFFECTS
Even among well-coupled donors and acceptors, substantial variations in CT
yields and rates have been observed. The fastest observed rates (subnanosecond) over
long-distances are for the RuII*/III/RhIII/II pair1,137 and for the oxidation of ZG by Et+* 120
and the reduction of RhIII by Et+*.121 Oxidation of guanine by Ap*,138 excited stilbene,122
or even by guanine radical after initial oxidation by photoexcited stilbene,139,140 is
substantially slower. To first order, this trend reflects the relative stacking of donors and
acceptors with the DNA duplex.
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In addition, some of these results may be reconciled in the context of considering
the effect of electrostatics on hole migration.113 This effect was directly demonstrated by
a study with RhIII* as the photooxidant wherein the position of a terminal phosphate was
varied.52 In this experiment, comparative damage at GG sites proximal and distal to the
Rh intercalation site was determined. Since the decomposition of the guanine cation
radical is slow, and the DNA between the GG sites was short and undamaged, the final
yield reflects both the relative extent of BET and the potentials at the GG sites. When a
phosphate anion is added to the end opposite to the rhodium, there is a small increase in
damage distribution towards the distal site. An extra phosphate anion on the same end as
the rhodium, however, leads to a several-fold change in relative damage. For one
sequence, relative damage at the proximal site increases from 16% to 56%. This argues
that local charge can have a strong effect, both on the rate of BET and on the rate of
migration. For AQ, which irreversibly injects a cation due to rapid oxygen quenching of
the triplet AQ radical anion, this effect is not present,141 consistent with a lack of BET,66
although given the low amount of distal damage in these constructs, it is not clear
whether a subtle change would be detectable.142 Importantly, in biological systems the
initial oxidation product is generally a guanine cation radical, without an anion radical
also being localized on the DNA. Hence, coulombic attraction will not inhibit transport
away from the injection site, as is the case for Ap* and Ap(-H)•, stilbene, AQ, and other
neutral photosensitizers.
26
1.2.7 ENERGETICS
The natural nucleosides of DNA are resistant to mild oxidation and reduction and
the radical cations and anions undergo secondary chemical reactions on the microsecond
time scale. Hence, the reversible potentials are not trivial to acquire.143 Approaches for
determining the nucleoside potentials fall into four categories: computational,
electrochemical, pulse radiolysis, and photooxidation studies (Table 1.1). A common
conclusion from all of these studies is that the oxidation potentials increase in the order
G<A<C~T.
Electrochemical measurements of base potentials are limited to organic solvents,
generally acetonitrile, DMSO, or DMF, due to the relative facile oxidation of water
versus the four bases. Considering the hydrophobic interior of DNA, potentials
determined under these conditions may be more relevant to DNA than potentials
determined in aqueous solution. To date, reversible electrochemistry has not been
achieved. Irreversible oxidation potentials of all four nucleosides have been measured,
which should be close to the standard potential.144 The values for dG are similar in
acetonitrile and DMSO,145 but substantially more positive and more negative values have
been found in chloroform145 and DMF,146 respectively. In chloroform, it was found that
the presence of dC, which allows the possibility of proton-coupled electron transfer
(PCET), lowers the oxidation peak of guanine by 340 mV.145
The closest that electrochemical experiments have come to measuring the
oxidation of guanine in its natural context in DNA are the electrocatalytic experiments
using mediators on indium tin oxide.147 That [Ru(bpy)3]3+ (E1/2 ~ 1.3 V; all potentials
27
Table 1.1. Experimental and Calculated Oxidation Potentials of Nucleotides
28
herein are vs. NHE) is a facile mediator for electrocatalytic oxidation of DNA indicates a
comparable potential for guanine.148 Analysis of oxidation rate using a variety of metal
complex mediators of different potentials supports this value. Notably, sufficiently high
ionic strength was used in these experiments to deconvolute the potential from the
affinity of the metal complex. This result was later validated by pulse radiolysis
experiments in DNA,149 where a potential of 1.22 ± 0.02 eV was found for guanine in
multiple sequence contexts. Although the absolute potentials from electrocatalysis are
approximate, they provide strong evidence for 5'-GG-3' being about 0.15 V lower in
potential than G with a 5'- pyrimidine.150
The latter result, that the 5'-G of GG doublets are lower in oxidation potential
versus G, has been extensively exploited in studies of the migration of charge in DNA,
where GG sites are used as shallow hole traps. Calculation and oxidative yield
experiments indicate that GGG acts as an even deeper well, although smaller differences
in potential between these sequences are found experimentally than by calculations,151
probably due to solvent interactions.152 Both calculation and experimental work support
preferential hole density on the 5'-G.99,153-156 More generally, there is strong correlation
between the calculated ab initio ionization potential of guanine in the different stacking
environments and the relative oxidative damage found between different guanines under
conditions that allow full equilibration of the injected charge.157 Stacking affects the
energies of all the bases, with the strongest perturbation due to the 5'-base.150,158
Vitally, all of these experiments probe the equilibrium potentials of the bases.
Random sequences of DNA have rugged potential landscapes, corresponding to extensive
29
static disorder. There are many conformational modes in DNA and its hydration layer
with time scales from picoseconds to seconds.127 As discussed in more detail below, the
energetics of the bases are coupled to these modes, introducing both static and dynamic
disorder to the system.
Given the challenge in properly coupling an individual photooxidant to DNA, it is
not surprising that few studies have attempted to determine rationally the driving force
dependence of CT. It has been established from several studies that CT does not occur
from an excited photooxidant hole donor to a higher energy hole acceptor. For example,
the metallointercalator [Rh(phi)2(bpy')]3+, tethered to DNA, can oxidize A, G, or C from
its excited state (E3+*/E2+ = 2.0 eV), but the metallointercalator [Ru(phen)(dppz)(bpy')]3+
(E3+/E2+ = 1.6 eV) oxidizes G, but not C.76 Similarly, [Ru(phen)(dppz)(bpy')]3+ and
ethidium bromide (E*+/E0 = 1.2V)121 are not competent to repair thymine dimers, but
photoexcited [Rh(phi)2(bpy')]3+ performs this repair, as does napthaldiimide (NDI)
(E*/E1- > 1.9 V).131,159 These studies include measurements with the hole donor tethered
far from the acceptor, with intervening low-potential double guanine sites, indicating that
even after charge is injected into DNA, there is some memory of the energy of its initial
state (Figure 1.6).
In support of this interpretation, CT can still occur far below the potential of the
DNA. In one example,160 an oxidized nitroxide (NO+/NO• ~ 1 V) was incorporated into a
duplex by covalent attachment to thymine. The [4Fe-4S] protein MutY (E3+/E2+ = 0.1 V)
was added, and the generation of the reduced nitroxide spin radical was observed by EPR
(yield ~ 50%). This chemistry was demonstrated to be DNA-mediated by two controls. A
30
Figure 1.6. Oxidation and repair of thymine dimer (~1.8 eV) by tethered photoexcited
[Rh(phi)2(bpy')]3+ (2.0 V) is unaffected by the intervening double guanine site (1.2 eV).
Oxidation of double guanine sites by [Ru(phen)(bpy')(dppz)]3+ (1.5 eV) is unaffected by
the presence of thymine dimer, which this oxidant lacks sufficient driving force to repair.
The latter result implies that guanine radical is not competent to repair thymine dimers, in
accordance with the known potentials. Hence either the guanine radical oxidized by
[Rh(phi)2(bpy')]3+ does not relax prior to migration to the thymine dimer, or the guanine
radical is not an intermediate in DNA-mediated oxidation of thymine dimer by
[Rh(phi)2(bpy')]3+.
31
noncleavable substrate analogue for MutY incorporated into DNA far from the label
increased the reduction yield, and partially saturating the electronic conjugation between
the label and the DNA substantially decreased the reduction yield.
A similar conundrum is involved in DNA-mediated CT in self-assembled
monolayers on gold (Figure 1.7).161 In these systems, a DNA monolayer is covalently
self-assembled on an electrode, and CT through the DNA is measured with an
electrochemical probe. At an applied potential greater than 0.4 V, the DNA adopts an
upright conformation. Although elastic motions can bring the DNA in contact with the
surface,162 these conditions require high salt and fairly positive potentials. DNA
mediation has been established for our system by the methods discussed above, i.e.,
mismatch and binding event discrimination, probe conjugation, linker-length dependence,
and by differential redox potential between direct contact and DNA-mediated
reduction.31,163-165 The window for these experiments on Au is limited by the potential of
Au-S reduction, about -0.5 V. DNA-mediated CT has been observed for a dozen different
probes spanning a full volt below this value.91,163-166 Importantly, the reductive limit is
600 mV below the reduction potential of cytosine.
Photooxidant-bridged DNA hairpins were employed to measure systematically
the dependence of CT rate on driving force.167 In these experiments, five photooxidants,
stilbene-4,4’-dicarboxamide (Sa, E*/- = 1.68 V), naphthalene-2,6-dicarboxamide (E*/- =
1.74 V), diphenylacetylene-4,4’-dicarboxamide (PA, E*/- = 2.02 V), NDI (E*/- = 2.93 V),
and phenanthrene-2,7-dicarboxamide (E*/- = 1.43 V), were employed as hole donors.
Guanosine, inosine, deazaguanosine, and 8-oxoguanosine were used as the hole
32
Figure 1.7. Scale diagram describing the relevant potentials for DNA-mediated CT
through DNA self-assembled monolayers on gold. The potentials of the individual
nucleotides are not accessible within the window of electrochemistry of DNA
monolayers on Au. Nevertheless, facile DNA-mediated electrochemistry is observed for
redox probes over DNA bridges. For all probes and sequences of well-matched duplexes,
the tunneling through the alkane linker is rate limiting (~30 s-1). Shown, in order from
top, are daunomycin, methylene blue, Redmond Red, and a [4Fe-4S] cluster similar to
those in the redox-active repair proteins EndoIII and MutY.
33
acceptors, either immediately adjacent to the bridging dye, or separated by a bridge of
two A-T base pairs. The measured charge separation and recombination rates fit well to
the Marcus-Levich-Jortner equation, with reorganization energies of 1.2 eV and 1.3 eV
respectively (Figure 1.8). The agreement is improved if a molecular based model for the
solvent is used and the Q-model is employed for the energy surfaces; in this case higher
reorganization energies are found.168 Further experiments varied the bridge length for the
St and PA duplexes, and found that the distance dependence increased for greater donor-
bridge energy separation.169
1.2.8. PROTON-COUPLED ELECTRON TRANSFER
If a participant in electron transfer is also an acid or base, the reduced and
oxidized species will often have different pKA values as well. In this case, it is likely that
CT will be coupled to proton transfer.170 Proton-coupled electron transfer (PCET) may be
more the rule than the exception in biological systems. Most redox-active groups in
biology are also subject to protonation or deprotonation, with the pKA dependent on the
redox state. Since complete proton transfer is unnecessary for substantial effects on the
coupled electron transfer rate, the question is not whether the electron transfer is proton-
coupled, but whether the coupling is significant so that proton transfer becomes rate-
limiting. In the case of Class I E. coli ribonucleotide reductase, PCET has been found to
occur over multiple steps.171-173
Each nucleotide in the double strand participates in stable hydrogen bonding with
its complement in a base pair. Hence, it is not surprising that CT between nucleotides
34
Figure 1.8. The driving force dependence for CT in photooxidant-bridged DNA hairpins
is determined from time-resolved transient absorption studies of a series of five stilbene-
derived photooxidants and four hole acceptor bases, following both charge separation
(filled) and charge recombination (empty). Case I,II (circles): donor and acceptor are in
contact. Case III,IV (triangles): donor and acceptor are separated by two TA base pairs.
Case I,III are fit only to charge separation rates (dotted), while Case II,IV are fit to both
charge separation and charge recombination rates (solid). Similar reorganization energies,
about 1 eV for the nuclear reorganization energy and ~0.2 V for the solvent
reorganization energy, are found for both Case II and Case IV. Adapted from Reference
167.
35
should be proton-coupled, although likely in a way that cannot be probed by the usual
assay of pH dependence, given that the base-pair protons are excluded from solvent. It
has been shown that oxidation of the aqueous, isolated nucleosides by Ap* is not proton-
coupled,174 but that does not exclude the possibility of PCET in the context of protons in
the base pair. Theoretical work predicts that double proton transfer between guanine and
cytosine lowers the guanine potential.175 Indeed, the cytosine radical has been directly
observed by transient absorption spectroscopy, after oxidation of DNA by SeO3•- and
SO4•- ions generated by pulse radiolysis.176 This might not be general, though, as the
mechanism of G-C oxidation in pulse radiolysis is strongly dependent on the chemical
interaction with the oxidizing radical.149,177 Similar evidence for PCET reduction of
thymidine base-paired to adenine has also been found.178
Furthermore, the pKA of the guanine cation radical has been measured to be 4,
near that of cytosine (4.5),179 and the neutral guanine radical is observed by nanosecond
transient absorption and EPR after oxidation by intercalated -[Ru(phen)2(dppz)]3+,180,181
supporting deprotonation of the guanine cation radical, presumably to the paired cytosine,
on a faster time scale. For DNA ionized by -irradiation at 77 K, ESR measurements find
that the equilibrium strongly favors the neutral guanine radical over the cation radical;
this held for guanine stacked between cytidine and for each guanine of the GGG
triplet.155 Although this evidence strongly supports proton transfer, it does not establish
coherent PCET, as proton transfer consequent to oxidation has been shown to be
favorable.182
36
Isotope experiments, which would establish whether proton transfer is rate-
limiting, have not been straightforward. Certainly, for oxidation by SO4•- ions, the charge
injection yield is decreased in D2O.177 In one experiment, deuterium replacement of acid
protons led to a three-fold decrease in the relative yield of damage of distal GGG to
proximal G sites.87 In another, CT between Ap(-H)• and G was found to exhibit a small
differential between D2O and H2O, consistent with PCET.183 A similar small differential
was observed in some sequences, but not in others, for CT between photoexcited NI and
Ptz.184 However, in the fluorescence experiments, the substitution of D2O for H2O also
affects excited state lifetimes.
At first, it might seem that the facile oxidation of CPC in competition with CPG
supports a PCET model.73 However, CPC oxidation is increased by base-pairing with
inosine, a high potential guanine analogue, indicating that PCET is not the mechanism of
CPC oxidation. Instead, this mechanism is enticingly similar to the proposed mechanism
for excited state relaxation in GC base pairs,185,186 which involves proton-coupled
exciplex formation and has also received experimental support,187,188 although it appears
that guanine-guanine stacking might prevent this relaxation.189 Based on this accumulated
evidence, it seems likely that PCET is involved in at least some charge injections to
guanine, and that neutral guanine radical is the persistent form of injected radical.
1.2.9. CHARACTERISTICS OF DNA CT
It is apparent that DNA mediates CT over long distances, and that the rate and
yield are sensitive to both the donor and acceptor identities and to the integrity of the
37
intervening -stack. Structural distortion of the DNA, or poor coupling of the donor or
acceptor to the DNA, sharply attenuate long-range CT. Furthermore, rapid CT is
conformationally gated, and the equilibrium conformation is not necessarily the CT-
active conformation.
1.3. DNA-MEDIATED CT MECHANISMS
Tautologically, all mechanisms of charge transport incorporate an electron
moving from a donor orbital to an acceptor orbital. The variation consists in the
identification of orbitals that mediate this transition, and the pathways that are coupled to
it. In a large biomolecule, such as DNA, complexity arises from the sheer number of
atoms involved. In this section, we will evaluate postulated mechanisms of long-range
CT in DNA in the context of the properties discussed above.
1.3.1. TRANSPORT THROUGH WATER, IONS, PHOSPHATES
An obvious source of conductivity in DNA is the highly charged phosphate
backbone. Indeed, one of the earliest models of CT through DNA involved transport
through the phosphates.190 A recent measurement of delocalization of a hole produced on
a single phosphate lends some credence to this model,191 although it is unclear whether
this delocalization can be transduced into conduction, and comparable measurements
have not observed this delocalization.192 In the phosphate conduction model, phosphates
on the edge of the DNA are directly ionized, and the hole rapidly hops through
isoenergetic phosphates. For this to occur, coupling between the phosphates must be
38
substantial. Even more importantly, oxidative damage must preferentially occur at
phosphates versus the base stack. Some calculations found that this was the case,193 but
later work demonstrated that this was due to neglecting the presence of water and
counterions that can shield the phosphate group’s negative charge.194 Theoretical and
experimental work suggests that photoionization is initiated at bases and not at
phosphates195 and that the energies of the ions, phosphates and sugar states are far from
the Fermi energy.196
Alternatively, the motions of water and ions can lead to apparent conduction.
DNA adsorbs several layers of high dielectric water,42 a primary condensation layer of
cations, and a secondary layer of condensed anions. Even under relatively dry conditions,
water and cations are still adsorbed. This layer plays a major role in the conformational
dynamics of DNA and mediates molecular recognition events with other biomolecules. In
particular, it seems certain that early conduction measurements were measuring ionic
conduction along the DNA, rather than properties of the DNA molecule itself.43
Ultimately, however, it is difficult to rationalize these models with the marked
sensitivity of long-range DNA-mediated CT to the integrity of stacking, as described in
Section 1.2. In contrast, changing the pH, ionic strength, or the identity of the salt has at
best a minor effect on CT, as long as well-coupled donors and acceptors are employed.
Even the removal of a phosphate along the bridge does not cause a measurable difference
in CT yield.77,78 Adding extra intervening phosphates, via the construction of triplex
DNA, actually lowers the competence for CT.197 Hence, it is apparent that DNA CT must
proceed through the base pairs, in the interior of the duplex.
39
1.3.2. SUPEREXCHANGE
Any medium is a superior pathway for charge transport in comparison to vacuum.
Superexchange is coherent orbital-mediated tunneling, where, for electron (hole)
transport, the high-energy LUMOs (HOMOs) on the pathway are virtually occupied,
allowing a probability and corresponding rate of transmission from the donor to the
acceptor. Following the Born-Oppenheimer approximation, the rate of superexchange can
be separated into the nuclear factor, n, and an electronic factor, e:
where
and
and G is the driving force, H0DA is the donor-acceptor coupling extrapolated to zero
bridge length, is a decay parameter characteristic of the bridge, and d is the bridge
length. The nuclear factor is a function solely of the identities and the environment of the
donor and acceptor. The electronic factor represents the electronic coupling between the
donor and acceptor, mediated by the bridge states. In the adiabatic limit, electronic
coupling is sufficiently strong such that the nuclear motion will determine the rate of
charge transfer. In the non-adiabatic limit, the electronic coupling is sufficiently weak
such that the electronic transition probability is less than unity at the transition state.9
40
Hence, long-range (greater than 1 nm) CT systems are generally treated as non-adiabatic,
though it has increasingly been recognized that changing the structure of even long
bridges can have a non-trivial affect on both G and .7,168,185,198 Ignoring this effect, the
only dependence of the rate on donor-acceptor distance is the exponential decay of the
donor-acceptor coupling with d, the length of the bridge, characterized by the parameter
. It is important to note that is generally not what is directly measured in experimental
systems. For systems that measure the yield of irreversible chemical products, competing
processes such as BET or equilibration will inevitably convolute with the inherent rate of
charge separation. Even for very fast charge traps, or spectroscopic based measurements
that can directly measure kCT, the exponential drop-off will not necessarily correspond to
if the nuclear factor is itself distance dependent.199 This restriction can be mitigated for
long-range CT, where the iterative changes in the bridge length are unlikely to affect G
and , but is significant for short-range CT.7
Furthermore, it is important to note that calculation of the CT rate requires precise
knowledge of the intervening electronic structure, which in turn is dependent on
molecular structure. If the mechanism or pathway changes with an increase in bridge
length, then the distance dependence will not be well represented by the electronic factor
. Also, conformational dynamics can lead to a time-dependent rate. If the equilibrium
structure is the best-coupled structure, dynamics will decrease the apparent CT rate.
Another important consideration is that is not independent of the bridge and
donor energies. For increasing difference between the donor and bridge energies,
increases according to:
41
where a is the intersite separation, V is the intersite coupling and is the donor-bridge
energy separation. As this separation decreases to below V, direct injection will
successfully compete with tunneling. Hence, if tunneling is occurring, is limited to
about 0.3 Å-1 (numerical calculations that properly treat the bridge as finite find about
0.2 Å-1).200 This supports the assignment of extremely shallow distance dependences to
incoherent processes; at least it excludes superexchange mediated by orbitals on the
individual bases of DNA. This model was supported by experiments in photooxidant
capped adenine tract hairpins.169 The oxidations of guanine by photoexcited Sa and of ZG
by photoexcited phenanthrene-2,7-dicarboxamide are of similar driving force, but the
latter pair are 0.25 eV lower in potential than the former pair. For each pair, the rate
constants were measured for varying lengths of an intervening adenine tract, and the
distance dependence was greater for the PA-ZG pair.
Superexchange has been most thoroughly characterized as a mechanism for CT
within and between redox-active proteins; charge-transfer reactions among proteins are
essential to all organisms. To a rough approximation, proteins can be treated as a
homogeneous medium with a single characteristic of 0.9 Å-1.201 The scatter for
individual proteins, however, spans several orders of magnitude, indicating that the
electronic structure and pathways vary strongly with the identity of the protein, and the
location of the donor and acceptor.202 For some pathways in proteins, conformational
dynamics have been shown to play an important role in dictating which pathways are
42
available.203 It is clear that proteins optimize charge transport not only by controlling
donor-acceptor distance and driving force, but by allowing a specific pathway, or
combination of pathways, to be available for superexchange. An essential lesson from
superexchange in proteins is that the most facile pathways determine the overall rate and
yield. Although DNA might appear to be a simple one-dimensional system, owing to the
extensive -stacking, experiments suggest a more complicated system.
1.3.2.1. COUPLING CONSTANTS IN DNA
No model of superexchange can be properly constructed without first considering
appropriate values for the coupling constants along the bridge.204 Given the structural
complexity, non-trivial assumptions are necessary to allow tractable calculations, each of
which have certain disadvantages. Furthermore, the stacking interaction of bases is a
particularly challenging one to computationally describe.205 It is important to consider
these couplings when developing a theoretical model. For example, a two-stranded
model206 for coherent DNA-mediated CT was recently published to fit the results of
work207 by Giese and coworkers. The model was only able to fit the data by taking
intrastrand AA coupling to be 0.52 eV, nearly an order of magnitude greater than the
time-averaged value found in typical calculations.208,209 The average couplings appear to
be about 80 meV for intrastrand GG, with somewhat lower intrastrand coupling between
AA, and smaller values for interstrand purine-purine couplings.
Increasingly, it has been clear that couplings are highly dependent upon the
geometry of the stacked bases. An early demonstration of this concept was the calculation
43
of coupling constants between base pairs for coordinates drawn for a large family of
crystallized duplexes.210 Even though this measurement was only for coordinates from a
set of crystals, each of which presumably corresponds to an equilibrium conformation,
variations in couplings were on the order of half of the values. In addition to this static
disorder, DNA is subject to extensive dynamic disorder on a full spectrum of time-scales.
A later study considered fluctuations from equilibrium conformations, using MD
simulations to access the transient structures (Figure 1.9).211 This study found even larger
variations in coupling, and found that HDA for GAAAAG varied by more than an order of
magnitude over the course of the 40 ps simulation. Interestingly, they also found that
transverse base motions, which affect stacking, are more significant than longitudinal
motions; this is consistent with recent work that found that shear, twist, and stretching
within base-pairs also affects coupling constants.212 They also found that peaks in
coupling over the bridge were more significant for GAAAAG than GTTTTG and nearly
absent in GATATG, in accordance with measured CT yields. Since then, similarly large
fluctuation-dependent variations have been found using a variety of computational
approaches,49,208,213,214 with fluctuations being most significant on the picosecond time-
scale.213 These studies have demonstrated that conformation also has a profound effect on
the transient nucleobase energies, as does solvent polarization.215 Interestingly,
calculations indicated that base energies tend to be correlated in the duplex,182 although
the relative ordering of base energies is preserved.208 These results offer a natural
explanation for the conformational gating that has been observed in long-range systems.
44
Figure 1.9. The time-dependent couplings between guanines separated by three different
four-base sequence contexts, based on conformations generated through molecular
dynamics. It is clear that the average value of coupling can be several orders of
magnitude lower than the maximum coupling. For the poorly stacked, flexible ATAT
sequence, strong coupling between the guanines is not achieved over the time-scale of the
simulation. Adapted from Reference 211.
45
1.3.2.2 REORGANIZATION ENERGY
Given the excellent correlation of theory and experiment for the driving-force
dependence of CT in stilbene-capped hairpins,167 the reorganization energy for those
systems is certainly close to 1 eV. For systems where the donor and acceptor are internal
to the -stack, as with intercalators, it is less clear, as these sites are far less solvent
exposed than the end-capped agents, such as Sa, Ptz, AQ, and napthalimide (NI).
Even for transfers between sites in DNA, the reorganization energy can vary
substantially. Sequence context, which affects couplings and site energies, is expected to
affect reorganization energy as well. Delocalization among multiple bases, which
decreases the effective amount of charge that must be transferred, lowers the
reorganization energy.216 One study found, by sampling many molecular dynamics
configurations of oligopurine•oligopyrimidine DNA, reorganization energy for nearest
neighbor hops to be about 1.1 eV for both adenine to adenine and for guanine to
guanine.49 Another study estimated 0.5 eV for adenine to adenine based on the spectral
density of intercalated ethidium bromide.217
It has been found that for short-range CT in DNA, changes in bridge length can
induce substantial changes in the reorganization energies.7,118 This finding is consistent
with Marcus’ classical description of the solvent reorganization energy:
where q is the change in charge, aD and aA are the donor and acceptor radii respectively,
and op and st are the optical and static dielectric constants, explicitly depends on the
46
donor-acceptor separation.9 An indirect length dependence of will also be incorporated
through the effect of molecular structure on the dielectric.
1.3.2.3. SUPEREXCHANGE IN DNA
Long-range charge transport over more than 50 Å seems incompatible with
superexchange, given its inherently strong distance dependence. Even a of 0.1 Å-1
implies a loss of over eight orders of magnitude in rate over 200 Å. However, most long-
range measurements either neglect yield122,159 or measure products formed on long time
scales.3 In the former case, long-range transport can reflect a small yield, while in the
latter case, products might be formed only on the time scale of milliseconds to seconds.
Fluorescence quenching of photoexcited 2-aminopurine by guanine 35 Å away54 has
shown that long-range CT can occur on a time scale that is defined by the nanosecond
lifetime of the 2-aminopurine excited state. Furthermore, the distance-independent
decomposition of CPA by photoexcited [Rh(phi)2(bpy')]3+ over 40 Å17 (Figure 1.10)
demonstrates that CT occurs at high yield at least as fast as BET between oxidized
adenine and [Rh(phi)2(bpy')]2+; the energetically similar reduction of [Rh(phi)2(phen')]3+
by [Ru(phen')2dppz]2+ over 41 Å is faster than 3 nanoseconds.1 Hence, it is clear that
DNA CT can occur over long distances on relatively short time scales, and any model
must account for this. For these reasons, superexchange models are not satisfactory for
DNA-mediated CT over long distances.
47
Figure 1.10. CT from photoexcited [Rh(phi)2(bpy')]3+ to N-cyclopropyladenosine (CPA)
across an adenine tract is distance-independent over 14 adenines. The rate of CT across
the adenine tract, then, must be much faster than BET from the first adenine to the
reduced rhodium. The driving force for recombination is only about 1.7 V, implying that
BET should not be in the inverted region, consistent with evidence that BET from
adenine to this rhodium complex is facile.68 The lack of distance-dependence, in a system
with a rapid competing process in BET and a charge trap that samples pre-equilibrium
CT dynamics, implies extensive delocalization across the bridge. Adapted from
Reference 17.
48
1.3.3. LOCALIZED HOPPING
The apparent contrast between theory and experiment led to extraordinary efforts
to challenge the validity of the measured CT rates and yields. Hopping models offer an
alternative that does not require exceptional coupling between bridge sites. Hopping, a
type of diffusive, incoherent transport, is the concatenation of multiple superexchange
steps, or “hops”, in which charge occupies the bridge between each hop. Hopping has
been proven as a mechanism in both natural171 and synthetic218 protein models. The
distance dependence of hopping is geometric, and hence shallower than the distance
dependence of superexchange.202,219 The reason for the shallower distance dependence is
intuitive; long, slow, superexchange steps are avoided.
1.3.3.1. NEAREST-NEIGHBOR MODELS
Although formal ballistic models do not distinguish superexchange from hopping,
it is most straightforward to treat hopping as a multi-step process. In this case, an injected
charge resides on the lowest potential base, guanine. This charge can diffusively migrate
along the DNA, mediated by short single-step superexchange with neighboring
guanines.219 The rate of a hop will depend on the distance and sequence context. The
fastest hop will be to neighboring guanine; hops through other nucleotides (e.g. GAG,
GCG, or GTG) will be slower. For the case of GCG, hopping is also allowed to the
guanine on the complementary strand (G+CG/CGC GCG/CG+C GCG+/CGC).
The most impressive evidence in support of this model is a series of
photooxidation experiments by the Majima group, and a series of STM conduction
49
experiments performed by the Tao group.45 Using an elegant experimental setup,220 they
form unambiguously covalent contacts to single DNA molecules under aqueous
conditions. They found a geometric dependence of the conductance on length for (GC)n
sequences, but insertion of an (AT)m sequence into the (GC)n sequence led to an
exponential decrease of the conductance with the length of the (AT)m sequence, as
predicted by the hopping model. However, they did not investigate sequences that
allowed purine-purine stacking, and were unable to find evidence for thermal
activation,221 although this was possibly due to the limited window of temperatures and
potentials that allowed device stability. Calculations on averaged structures confirm that
the alternating purine-pyrimidine sequence attenuates delocalization for this system and
reproduce the data well.222 Furthermore, more recent experiments using the same
system223 and a similar approach using a mechanical break junction224 found a much
smaller effect on conduction from increasing AT content. This was ascribed to the latter
experiments being performed with DNA that was more likely to form the B-conformation
versus the (GC)n sequences. For DNA covalently bridging a carbon nanotube gap, there
appears to be no sequence dependence when the GC content of random sequence DNA is
varied.225
Majima and colleagues have used transient absorption to study the oxidation of
Ptz by photoexcited NI, linked by varying sequences of DNA.4 The sequences near the
hole acceptors and donors were kept constant, and a central region varied with (GA)n or
(GT)n; complementary studies separated the donor and acceptor by only adenines.226,227
Each system fits well to a geometric dependence of rate on bridge length. They found the
50
rate of hopping to be 2x1010 s-1 for adenine to adenine hopping, 7.6x107 s-1 for G+-A-G
G-A-G+ hopping, and about 2x105 s-1 for G+-T-G G-T-G+ hopping. Although this is
strong evidence for multistep hopping in these systems, they have noted that it does not
necessarily require that the intermediate states be completely localized on individual
nucleotides.227 More generally, the researchers were able to distinguish between hole
injection and hole arrival, showing that the two are not coincident over long distances.
Similar results have been observed for CT between DNA-capping stilbenes, with the
transition between single and multistep CT at about two intervening AT base pairs.125,126
1.3.3.2. THERMALLY INDUCED HOPPING
Although hopping between guanine sites can explain many features of the
propagation of holes in mixed sequences, it is not sufficient to explain facile charge
transport through adenine tracts.228 Occupation of adenines during CT has been
demonstrated both by a direct chemical probe73 and by the observation of facile and
weakly distance-dependent transport of holes across long adenine bridges,17,31,207,229-231
even when BET competes with equilibration.31 This can be explained by a reasonable
modification of the hopping model, whereby a hole on guanine can be thermally excited
to occupy an adenine tract.228,232,233 Although this will be disfavored in the case of mixed-
sequence DNA in preference to guanine hopping, it can be much more favorable than the
slow hop through a long AT sequence. For isoenergetic adenines, this model does not
sufficiently explain the distance independence,234 but if the adenines on the edge of the
tract are higher in potential than the interior adenines, then the apparent yield of CT will
51
be distance-independent.235 This explanation, although reasonable for (A)n bridges on the
strand complementary to the guanine sites, does not support the shallow distance
dependence observed when the (A)n bridge is in the same strand,229,236 for which the edge
adenines should be of lower energy versus internal adenines. It would be interesting to
determine what effect incorporating the stacking-dependent adenine energetics would
have on the theoretical predictions of the thermally induced hopping model.228
The most compelling evidence for thermal activation comes from a biochemical
trapping assay of G+/An/GGG, where the yield of GGG versus G damage was quantified
after hole injection from a sugar radical near the single G site.207 A steep distance
dependence for n 3 was followed by a flat distance dependence for n 3. This is
consistent with two mechanisms at play, where the steep distance dependence
corresponds to CT through superexchange across the AT bridge, and the flat regime is
where superexchange is sufficiently slow for thermally induced hopping across the
adenine tract to become the dominant mechanism. It is important to note, however, that
this dependence looks identical to that found for the rigid Et+ base-pair surrogate.27 In the
latter case, the dependence was caused not by a fundamental shift in mechanism, but
rather by the rate-limiting injection from the hole donor. Stilbene-capped hairpin
systems125 and AQ-capped duplexes236 show a similar, though much more gradual,
positive second-order change in the slope with distance. As shall be discussed in greater
detail below, delocalized mechanisms can also explain facile transport through adenine
tracts. Ultimately, a change in slope on its own is not sufficient to justify a crossover in
mechanism.237
52
1.3.3.3. VARIABLE-RANGE MODELS
All the mechanisms listed above can be considered together, as components of a
variable-range hopping model. Here, a hole is allowed to migrate by superexchange to
any other site, rather than being limited to nearest neighbors.238 The most probable sites
will be the closest low-potential sites, i.e. guanines. Hence, the hole will hop from
guanine to guanine through the DNA, preferring intrastrand transfer, but able to exploit
interstrand transfer or thermally induced hopping onto adenine tracts where the sequence
does not allow more favorable pathways. Even unfavorable pathways are possible,
although slow. Theoretical treatments using this model have been successful in modeling
some biochemical experiments,239 although it was demonstrated that introducing static
disorder substantially degrades the success of variable-range hopping models that rely on
localized states (Figure 1.11).217 In turn, dynamic disorder, analogous to the
conformational gating discussed above, can assist hopping in a rugged landscape.113
One challenge that is common to all localized hopping models is the explanation
of the mismatch discrimination that has been observed in nearly every system studied.
One proposal was that mismatches allow water access to preferentially quench CT at
guanine through proton abstraction or other chemical reaction.232 This model was
supported by the observation that GT mismatches affected distal yield more than AA
mismatches, and that methylation of the most acidic residue of a guanine opposite an
abasic site restores CT.87 This model has not stood up to more extensive measurements,
however, as AC mismatches attenuate CT more than GT mismatches. It has also been
shown that GT mismatches lower the yield of CT by lowering the rate,4 and GT
53
Figure 1.11. The variable-range hopping model predicts a shallow distance dependence
for the rate of CT between G and GGG across an adenine tract of the opposing strand.
Delocalized states, even in the absence of disorder (dashed lines), yield larger and
shallower CT rates due to the smaller reorganization energy, and a shorter effective
bridge length. In the presence of static disorder (solid lines), localized hopping is
substantially attenuated due to the rugged energy landscape. Delocalized hopping,
however, is relatively unaffected by static disorder, as the coupling is strong enough to
allow tunneling through local barriers. Adapted from Reference 217, which fit data of
Reference 207 using the interbase couplings of Reference 212, which are similar to the
most recent calculated values.208
54
mismatches attenuate CT even under applied potentials insufficient for guanine
oxidation.30 Alternatively, mismatched base pairs might have lower couplings to the
neighboring bases than matched pairs.62 Particularly, they are less stacked, and sample
more unstacked configurations. A similar argument can then be made to explain how
DNA-binding proteins that bend the -stack also attenuate CT.
A more profound problem with localized hopping models is the apparent
“memory” that a charge has of the energy of the state from which it was injected. The
intermediate in a localized hopping model is the cation or neutral radical on guanine or
adenine. Oxidation of cytidine or thymidine by these species is taken to be highly
unfavorable. Hence, the energy of injected charge should not affect the nature of the
intermediate over long distances. This is not consistent with the evidence from thymine
dimer159 or CPC75,76 oxidation, where oxidants competent for guanine oxidation, but not
pyrimidine oxidation, were unable to decompose these species over long distance.
Oxidants that are competent for thymine dimer repair or CPC decomposition, however,
remain competent to decompose these oxidation reporters even with intervening low-
energy guanines. For an extended (A)20 bridge separating Ptz and photoexcited NI,
central double guanine does attenuate CT yield by about half, indicating that over a very
long piece of DNA relaxation of the cation does occur.227
Localized hopping is also inconsistent with electrochemical measurements, where
the Fermi level is maintained up to a volt below the potentials of the bases.58,88,164,240,241
Consider as an example where the electrode is at the potential of an [4Fe-4S]-containing
protein (0.1 V), and injection is into cytidine (~ -1 V),144,174 the most readily reduced
55
base, for an unfavorable driving force of at least 1.0 V. Although the coupling between
cytidine and the metal is mediated by a long saturated linker, and hence will be small, let
the coupling correspond to a generous value for two stacked bases, HDA ~ 0.2 eV, and
take the reorganization energy as being 0.5 V. According to a simple nonadiabatic
Marcus-derived expression,116,242 the injection rate is no greater than 0.002 s-1; for
realistic values for the molecule-metal coupling this injection rate would necessarily be
far lower. This is slower than the linker-limited rate found through DNA of about
30 s-1.18,240 Effectively, this discrepancy reflects the inherent unfavorability of thermal
activation far from the bridge potential.
1.3.4. DELOCALIZED MECHANISMS
The models discussed so far each assume localization of a hole on a single base.
Although the couplings between bases might be expected to allow delocalization,
disorder in the bath should rapidly localize charges onto a single site as long as
reorganization energy is greater than interbase coupling. However, there is some
experimental evidence for delocalization of charges, such as the effect of stacking
interactions on the pKA of the adenine cation radical,243 or the competition of CPC with
CPG for oxidation.76 It has also been demonstrated that static disorder attenuates rapid
hopping by creating low potential bottlenecks.238 This can be alleviated by allowing
delocalization of the charge; in this case, static disorder is partially averaged.217,244 In
conjuction with the known role of conformational gating, the obvious candidate for the
delocalized state is the polaron.245,246
56
1.3.4.1. POLARON HOPPING AND GATING MECHANISMS
Whenever charge is injected into a molecule, the environment will polarize in
response, effectively partially delocalizing the charge and lowering the energy of the
system.247 Since the energy of the polaron is different than that of the purely localized
charge, the presence of a polaron will affect the CT behavior of the system, in a way
largely dependent on the polaron size. Much as PCET is inevitable for any charge-
transfer participant with acidic protons, polaron formation is inevitable whenever CT
proceeds with bridge occupation. The essential questions are whether the polarization
occurs on a time-scale that can impact the CT process, which relaxation modes will be
coupled to the polaron formation, and how much the polaron is stabilized relative to the
completely localized state.
At first order, polarization of the environment in response to charge injection does
not violate the tight-binding assumption. In this case, although DNA conformation, ion
distribution, and water orientation all restructure as a result of charge migration, the
effect on CT efficiency and rate will be via a change in the site energies on the bases, and
gated by the time-scale of environmental polarization. It is important to note that small
polaron formation slows charge migration, as the site energy is lowered, and hence the
activation energy of each hop is increased; this leads to dynamic disorder, distinct from
the static disorder discussed above. The exception is if the polaron can move by drift,
where the orbitals of the donor and acceptor states overlap, so that CT occurs in the
adiabatic limit. This results in transport that is faster than hopping, especially as it can be
57
activationless.247 However, drift is most rapid between isoenergetic sites, so it is not a
likely mechanism in the presence of static disorder, unless gated by conformational
fluctuation of site energies.
Any description of polarons must take into account the structural rearrangement
that provides the polarization. Lattice motion has been well treated in terms of
deformations along the hydrogen bonds between the base pairs.248-250 This treatment is
particularly instructive regarding the effect of increased coupling between the lattice
motion and the charge; high coupling implies a higher activation energy for individual
hops and a higher probability for trapping. Hence, thermal activation is taken as evidence
for small polaron trapping. There have been contradictory results on the temperature
dependence of CT in conductivity measurements,23,221 though photooxidation studies
have unambiguously shown an increase in long-range CT rate184 and yield54,68 with
temperature. Whether this temperature dependence is due to conformational gating, small
polaron activation, or activation of localized hopping is not immediately obvious.
Ultimately, the distinction between these cases is not sharp. Conformational dynamics
influence bridge energy, and hence the activation energy for polaron drift. Calculations
suggest that ion fluctuations, in particular, could sufficiently modulate the potential of a
bridging sequence of DNA to permit polaron equilibration between two sites.50 Given the
ambiguity in experiments where counterion identity and concentration have been
varied,51-54 water re-orientation is more likely than ion motion to gate polaron formation.
Sufficient polarization of the environment will lead to formation of a large
polaron. In this case, the polarization distortion extends far beyond the lattice site, i.e. the
58
individual base (Figure 1.12). Polarization over a large range must involve the medium,
water. Calculation has supported that these polarons form by water reorientation
delocalized over 2 to 5 base pairs, depending on the sequence.251-252 Large polaron
formation can have both positive and negative consequences for CT: self-trapping can
decrease the rate of individual hopping steps, as is the case for small polarons, but
delocalization decreases the distance between individual steps. Furthermore, for periodic
sequences, drift can substantially increase the rate of individual hops, by lowering the
activation energy for incoherent transport.
Critically, polaron drift can explain important features of DNA-mediated CT as
discussed in the previous sections. As discussed above, the observed dependences of CT
rates and yield on distance across adenine tracts is too shallow to be readily reconciled
with thermal activation and localized hopping.17,125,207,236 Rapid polaron drift across
adenine tracts, in concert with inhibition of BET from adenine to guanine due to polaron
self-trapping, has been predicted to provide a shallow distance dependence.253
Furthermore, since the calculated polaron size is ~ 4 adenines, the steep distance
dependence that has been found for tracts shorter than this length naturally corresponds to
these sequences not supporting polaron formation.
This mechanism is not limited to adenine tracts, as polaron formation is predicted
over mixed polypurine sites, with significant population of high-energy
pyrimidines.252,254 This is consistent with the oxidative damage observed at the fast trap
CPC despite the presence of guanine75 and the preferential damage at thymine in
constructs containing only thymine or adenine.69 As described above, the resulting
59
Figure 1.12. Formation of a large polaron. Upon charge injection, a hole is initially
localized on a single base (red). Reorientation of the environment, including neighboring
bases and the hydration layer, lowers the energy of the hole. Delocalization occurs to the
extent that the coupling between the bases balances the unfavorable decrease in the
reorganization energy.
60
delocalization serves as a mechanism for dynamic motions, that allow polaron formation,
to alleviate the barrier to CT generated by the static disorder of site energies in DNA, and
is consistent with the observed long-range migration of CT through mixed DNA
sequences.3,103 In this model, the effective hopping rates observed by the Majima group
could correspond to hops between delocalized polaron sites.4,226,227
Physical identification of the polarization medium allows calculation of the
polaron properties, particularly the speed limit on polaron migration imposed by the rate
of repolarization. For drift along an adenine tract, water reorientation limits polaron
mobility to about 3x10-3 cm2/(V s).255 This mobility can be related to the conductivity of
a single DNA between two carbon nanotubes,34 where a mixed, aperiodic sequence
should decrease the mobility of a polaron. Here, a resistance of about 3 M was
observed in a fifteen base-pair duplex. Although the number of charge carriers was
unknown, it certainly cannot be less than unity or greater than fifteen, the number of base
pairs. Within that range, the mobility is constrained to between 3 x 10-2 cm2/(V sec) and 5
x 10-1 cm2/(V sec). It will be interesting if theoretical evaluation is able to rationalize
these values, as they appear inconsistent with polaron drift.
1.3.4.2. DOMAIN DELOCALIZATION
Evidence for delocalization in DNA has come from recent insights into long-lived
excited states and exciplexes. It is well known that the individual nucleotides of DNA
rapidly relax upon excitation, with a low fluorescence quantum yield.256 This property is
61
essential for the molecule of life, as long-lived excited states would render the genetic
material prone to photodamage. For mixed sequence DNA as well, relaxation is rapid
(subpicosecond to picosecond), but in a manner highly dependent on the specific
sequence context.257 Recently, however, femtosecond studies of purine repeats in both
oligonucleotides and duplexes have found much longer lifetimes for ground state
recovery,189 possibly due to exciton or exciplex states. Critically, for B-form DNA, it is
base stacking rather than base-pairing interactions that are most critical in achieving long-
lived states.258 The extent of delocalization is still under debate. Although some states
might extend over 4 to 8 base pairs,259,260 others are delocalized over only two base
pairs,261 particularly in single strands. There is some computational support for this
delocalization, as well. DFT calculations find that eximers can delocalize over several
adenines,262 and it has been found that fluctuations in the on-site energies of neighboring
bases is highly correlated.214 Recent calculations that incorporate static and dynamic
disorder and solvent effects have shown that such transient delocalization can occur over
several base pairs.263
Despite experimental and computational support for delocalization, as described
above, there are profound theoretical arguments against delocalization models in DNA.
A variety of computational studies have found that solvent and ion motions will strongly
localize injected charge to a single or only a few nucleotides.152,264,265 Importantly, these
studies have mostly been limited to considerations of equilibrated or averaged structures.
It will be important to determine whether solvent-induced localization is maintained in
the presence of dynamic disorder.
62
A recent computational study266 on the dynamics of electric fields produced by
DNA, water, and ions was able to reproduce time-resolved Stokes shift data that directly
measures those dynamics.127 This study found no evidence of subpopulations where the
DNA had particular electric fields beyond the Gaussian distribution. However, the
Gaussian tails could in principle be the very states that mediate CT. All the data suggest
that delocalization must be highly transient, and over very long distances, a substantial
amount of incoherent hopping will also occur.
Perhaps the most distinctive feature of DNA-mediated CT that has been observed
in recent years is the periodic length dependence of CT yield across adenine tracts for
some systems (Figure 1.13). This dependence was clear, with the same period of 3 to 4
base pairs, for coherent transport from Ap* to guanine54 and for total CT from
[Rh(phi)2(bpy')]*3+ to CPG, as will be discussed in Chapter 2.68 This periodicity was
shown to be with respect to adenine tract length, rather than with respect to donor-
acceptor distance; by measuring the decomposition of CPA moved serially along an
adenine tract of constant length, no periodicity is observed.17 For CT from photoexcited
AQ to guanine, the periodicity is less apparent, but this is likely due to the quenching of
the radical anion of AQ by oxygen, which allows charge equilibration. Interestingly, Ap*
oxidation of CPG across adenine tracts is smoothly monotonic, but separating the Ap*
from the adenine tract with three inosines restores the periodicity. Clearly, the rapid BET
associated with Ap* allows duplexes that are well suited to forward transport also to
better mediate BET, suppressing the periodicity. It should be noted that the inosine tract
is a high potential barrier to oxidation by Ap.120 It lowers both forward CT and BET, but
63
Figure 1.13. Equivalent periodicities with the same period and temperature dependence
are observed for (B) the single-step oxidation of guanine by Ap* and (A) the total
oxidation of CPG by photoexcited [Rh(phi)2(bpy')]3+. Temperature increases from purple
to red. Errors are given in (A) as 90% SEM.54,68 Adapted from References 54 and 68.
64
since the former competes with the nanosecond Ap* fluorescence lifetime, and the latter
competes with picosecond ring opening, BET should be comparatively more attenuated.
With BET suppressed, periodicity is again apparent.
A periodic A-tract dependence indicates that some adenine tract lengths mediate
CT superior to others. Based on our experiments, this length is about three or four base
pairs. In light of the extensive evidence for delocalization cited above, we characterize
this CT-active tract as a delocalized domain. The role of conformational gating, then, is
to generate this CT-active state. An adenine tract length that allows an integer number of
these states allows facile CT; transport across other tracts requires dephasing processes,
such as drift or hopping. Because these domains are, by their nature, transient, these
effects will only be seen in experiments where the donor and acceptor are well-coupled to
the bridge, and where injection and arrival can be observed on a fast time scale,
decoupled from other pathways, such as BET. Critically, domain delocalization readily
explains the facile competition between CPC and CPG,74 and the ability of DNA to mediate
CT far below the base potentials.161,164
1.4. SUMMARY
It is clear that DNA, when adequately coupled between the donor and acceptor,
can competently mediate CT over long distances. This property is dependent on, and
hence diagnostic of, the integrity of base stacking. Furthermore, long-range DNA-
mediated CT is thermally activated in a manner dependent on the dynamical stacking of
the bridge, indicating that conformational gating is convoluted with the CT rate.
65
Theoretically, CT over long molecular distances cannot be assigned to superexchange.
Incoherent transport must play a role, although evidence does support coherent transport
over at least 30 Å in some systems. Assigning the intermediates as guanine cation
radicals in the context of a variable-range hopping model is sufficient to explain some
gross features of DNA-mediated CT, but this model cannot explain long-range
coherence. Transient delocalization plays an important role, at least with some sequences.
Identifying the extent to which delocalization occurs, including via polaron formation,
will be particularly important for understanding DNA CT mediated at potentials below
those of the individual nucleotides.
Any model for DNA CT must consider the effects of static and dynamic disorder.
For most models, static disorder attenuates long-range CT. Since DNA has many sources
of static disorder in the site energies, inter-site couplings, and reorganization energies, it
is unlikely that calculations performed on uniform ideal structures with a single repeating
base pair will be relevant to understanding experimental results. On the other hand,
dynamic disorder has the potential to alleviate the challenge posed by static disorder, by
allowing transient structures to form with less rugged energetic landscapes. As long as
the equilibrium conformation is not the most CT-active conformation, this condition will
hold for most pathways, whether incoherent or coherent. Computational studies have
begun to appear that consider what CT-active states look like;267 it will be a challenge to
experimentalists to evaluate these exciting predictions.
CT between a donor and acceptor will always proceed through the fastest
pathways available. In a dynamic, structurally complex molecule like DNA, multiple
66
time scales describe the energetic and coupling landscapes, and hence there will be a
time-dependent ensemble of pathways. This ensemble is even larger when delocalized
states are allowed, whether they are transiently formed prior to, concurrently with, or
after charge injection. For conditions that deplete available pathways, whether through
rigidifying the duplex, disrupting donor and acceptor coupling to the bridge, or by
introducing structural distortion, slower CT and conduction will inevitably result.
In this context, correlating the distance dependence to the value of the electronic factor
of the CT rate equation requires a high level of experimental support. It is unlikely that
any of the measured distance dependences correspond to the distance dependence of the
purely electronic component of CT through DNA. Nevertheless, the effective distance
dependence over long distances compares favorably with common molecular wires such
as oligophenylenevinylene and oligophenyleneethynylene, indicating a promising role for
DNA in molecular electronics.
1.5. UNANSWERED QUESTIONS
It should be clear from this chapter that DNA-mediated CT does not pose a
challenge to the fundamental theories of electron and hole transport. Ultimately, charge-
transfer events only occur with the rates predicted by Marcus’ theory. For a molecule as
large and complicated as DNA, however, the parameters for the Marcus equation are not
trivial to determine. Each conformation of a given DNA offers many pathways, and the
extent of dynamical disorder can lead to the failure of the Condon approximation.
Furthermore, in the context of hopping and drift, the nature of the states that mediate
67
charge transport vary with sequence and sequence-dependent dynamics. What these
states are: localized radical cations, localized neutral radicals, large polarons, delocalized
domains, or a combination, will be different based on the properties of the specific donor,
DNA bridge, and acceptor. Understanding what conditions lead to what mechanism of
transport is important, as the physical nature of charge injection and migration in DNA
undoubtedly influences CT between DNA and redox-active DNA-binding
proteins,5,17,93,94 and the cellular defense against oxidizing radicals.105,268-271
Particular experiments that require more attention by theorists are the
electrochemical experiments in DNA films. In these experiments, the Fermi level is held
to potentials far from those of the bridge states, and yet many of the same properties are
observed here as are observed in solution and device experiments that are at profoundly
different energies. Insight into this process will undoubtedly also help elucidate DNA-
mediated CT in general.
Ultimately, single-molecule conductivity experiments have the most potential for
determining details of DNA-mediated CT, due to the strong control of driving force and
online measurement of current. The main challenges for these experiments is maintaining
the DNA in its native structure, and establishing that the observed current is, in fact, due
to the DNA. These can be easily determined by the proper choice of controls.
If the past fifteen years of DNA-mediated CT are any indication, the synergy
between the applications of DNA in devices and biology, and theoretical and
experimental efforts to elucidate the mechanism, will continue to advance both areas of
68
study. Certainly, bringing these different perspectives together offers both a challenge
and an opportunity.
69
1.6. REFERENCES
(1) Murphy, C.J.; Arkin, M.R.; Jenkins, Y.; Ghatlia, N.D.; Bossmann, S.H.; Turro, N.J.;
Barton, J.K. Science 1993, 262, 1025.
(2) Murphy, C.J.; Arkin, M.R.; Ghatlia, N.D.; Bossman, S.H.; Turro, N.J.; Barton, J.K.
Proc. Natl. Acad. Sci. U.S.A. 1994, 91, 5315.
(3) Nuñez, M.E.; Hall, D.B.; Barton, J.K. Chem. Biol. 1999, 6, 85.
(4) Takada, T.; Kawai, K.; Fujitsuka, M.; Majima, T. Proc. Natl. Acad. Sci. U.S.A. 2004,
101, 14002.
(5) Lee, P.E.; Demple, B.; Barton, J.K. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 13164.
(6) Osakada, Y.; Kawai, K.; Fujitsuka, M.; Majima, T. Proc. Natl. Acad. Sci. U.S.A.
2006, 103, 18072.
(7) Davis, W.B.; Hess, S.; Naydenova, I.; Haselsberger, R.; Ogrodnik, A.; Newton, M.D.;
Michel-Beyerle, M.-E. J. Am. Chem. Soc. 2002, 124, 2422.
(8) von Feilitszch, T.; Tuma, J.; Neubauer, H.; Verdier, L.; Haselsberger, R.; Feick, R.;
Gurzadyan, G.; Voityuk, A.A.; Griesinger, C.; Michel-Beyerle, M.E. J. Phys. Chem. B
2008, 112, 973.
(9) Marcus, R.A.; Sutin, N. Biochim. Biophys. Acta 1985, 811, 265.
(10) Turro, N.J.; Barton, J.K. J. Biol. Inorg. Chem. 1998, 3, 201.
(11) Endres, R.G.; Cox, D.L.; Singh, R.R.P. Rev. Mod. Phys. 2004, 76, 195.
(12) Chen, F.; Hihath, J.; Huang, Z.; Li, X.; Tao, N.J. Annu. Rev. Phys. Chem. 2007, 58,
535.
70
(13) Müller, K.-H. Phys. Rev. B 2006, 73, 045403.
(14) Kelley, S. O.; Barton, J. K. Science 1999, 283, 375.
(15) Delaney, S.; Pascaly, M.; Bhattacharya, P.K.; Han, K.; Barton, J.K. Inorg. Chem.
2002, 41, 1966.
(16) Tada, T.; Kondo, M.; Yoshizawa, K. ChemPhysChem 2003, 4, 1256.
(17) Augustyn, K.E.; Genereux, J.C.; Barton, J.K. Angew. Chem. Int. Ed. 2007, 46, 5731.
(18) Gorodetsky, A.A.; Green, O.; Yavin, E.; Barton, J.K. Bioconjugate Chem. 2007, 18,
1434.
(19) Fink, H.W.; Schoenberger, C. Nature 1999, 398, 407.
(20) de Pablo, P.J.; Moreno-Herrero, F.; Colchero, J.; Gómez Herrero, J.; Herrero, P.;
Baró, A.M.; Ordejón, P.; Soler, J.M.; Artacho, E. Phys. Rev. Lett. 2000, 85, 4992.
(21) Kasumov, A.Yu.; Kociak, M.; Guéron, S.; Reulet, B.; Volkov, V.T.; Klinov, D.V.;
Bouchiat, H. Science 2001 291 280.
(22) Storm, A.J.; van Noort, J.; de Vries, S.; Dekker, C. Appl. Phys. Lett. 2001, 79, 3881.
(23) Yoo, K.-H.; Ha, D.H.; Lee, J.-O.; Park, J.W.; Kim, J.; Kim, J.J.; Lee, H.-Y.; Kawai,
T.; Choi, H.Y. Phys. Rev. Lett. 2001, 87, 198102.
(24) Odom, D.T.; Dill, E.A.; Barton, J.K. Chem. Biol. 2000, 7, 475.
(25) Schuster, G. B., Ed. Long-Range Charge Transfer in DNA, I and II; Springer: New
York, 2004; Vols. 236, 237.
(26) Wagenknecht, H. A., Ed. Charge Transfer in DNA; Wiley-VCH: Weinheim, 2005.
(27) Valis, L.; Wang, Q.; Raytchev, M.; Buchvarov, I.; Wagenknecht, H.-A.; Fiebig, T.
Proc. Natl. Acad. Sci. U.S.A. 2006 103, 10192.
71
(28) Meade, T.J.; Kayyem, J.F. Angew. Chem. Int. Ed. 1995, 34, 352.
(29) Lyndon Pittman, T.; Miao, W. J. Phys. Chem. C 2008 112, 16999.
(30) Boon, E.M.; Jackson, N.M.; Wrightman, M.D.; Kelley, S.O.; Hill, M.G.; Barton,
J.K. J. Phys. Chem. B 2003, 107, 11805.
(31) Drummond, T.G.; Hill, M.G.; Barton, J.K. J. Am. Chem. Soc. 2004, 126, 15010.
(32) Watson, J.D.; Crick, F.H.C. Nature 1953, 171, 737.
(33) Watson, J.D.; Crick, F.H.C. Nature 1953, 171, 964.
(34) Guo, X.; Gorodetsky, A.A.; Hone, J.; Barton, J.K.; Nuckolls, C. Nat. Nanotech.
2008, 3, 163.
(35) O’Neill, M.A.; Becker, H.C.; Wan, C.; Barton, J.K.; Zewail, A.H. Angew. Chem. Int.
Ed. 2003, 42, 5896.
(36) Melvin, T.; Botchway, S.; Parker, A.W.; O’Neill, P. J. Chem. Soc., Chem. Commun.
1995, 6, 653.
(37) Cohen, H.; Nogues, C.; Naaman, R.; Porath, D. Proc. Natl. Acad. Sci. U.S.A. 2005,
102, 11589.
(38) Cohen, H.; Nogues, C.; Ullien, D.; Daube, S.; Naaman, R. ; Porath, D. Far. Disc.
2006, 131, 367.
(39) Ceres, D.M.; Barton, J.K. J. Am. Chem. Soc. 2003, 125, 14964.
(40) Boon, E. M.; Ceres, D. M.; Drummond, T. G.; Hill, M. G.; Barton J. K. Nat.
Biotech. 2000, 18, 1096.
(41) Calladine, C.R.; Drew, H.R.; Luisi, B.F.; Travers, A.A., Understanding DNA: The
Molecule and How it Works, Elsevier Academic Press: San Diego, 2004, 304.
72
(42) Pal, S.K.; Zewail, A.H. Chem. Rev. 2004, 104, 2099.
(43) Yamahata, C.; Collard, D.; Takekawa, T.; Kumemura, M.; Hashiguchi, G.; Fujita, H.
Biophys. J. 2008, 94, 63.
(44) Han Ha, D.; Nham, H.; Yoo, K.-H.; So, H.-m.; Lee, H.-Y.; Kawai, T. Chem. Phys.
Lett. 2002, 355, 405.
(45) Xu, B.; Zhang, P.; Li, X.; Tao, N. Nano Lett. 2004, 4, 1105.
(46) Nogues, C.; Cohen, S.R.; Daube, S.; Apter, N.; Naaman, R. J. Phys. Chem. B 2006,
110, 8910.
(47) Xiao, X.; Xu, B.; Tao, N.J. Nano Lett. 2004, 4, 267.
(48) Boon, E.M.; Barton, J.K. Bioconjugate Chem. 2003, 14, 1140.
(49) Steinbrecher, T.; Koslowski, T.; Case, D.A. J. Phys. Chem. B 2008, 112, 16935.
(50) Barnett, R. N.; Cleveland, C. L.; Joy, A.; Landman, U.; Schuster, G. B. Science
2001, 294, 567.
(51) Williams, T.T.; Odom, D.T.; Barton. J.K. J. Am. Chem. Soc. 2000, 122, 9048.
(52) Williams, T.T.; Barton, J.K. J. Am. Chem. Soc. 2002, 124, 1841.
(53) Douki, T.; Angelov, D.; Cadet, J. J. Am. Chem. Soc. 2001, 123, 11360.
(54) O’Neill, M.A.; Barton, J.K. J. Am. Chem. Soc. 2004, 126, 11471.
(55) Blaustein, G.S.; Demas, B.; Lewis, F.D.; Burin, A.L. J. Am. Chem. Soc. 2009, 131,
400.
(56) Abdou, I.M.; Sartor, V.; Cao, H.; Schuster, G.B. J. Am. Chem. Soc. 2001, 123, 6696.
(57) O’Neill, M.A.; Barton, J.K. J. Am. Chem. Soc. 2002, 124, 13053.
(58) Boon, E. M.; Salas, J. E.; Barton, J. K. Nat. Biotech. 2002, 20, 282.
73
(59) Okamoto, A.; Kamei, T.; Saito, I. J. Am. Chem. Soc. 2006, 128, 658.
(60) Kelley, S.O.; Barton, J.K. Chem. Biol. 1998, 5, 413.
(61) Bhattacharya, P. K.; Barton, J. K., J. Am. Chem. Soc. 2001, 123, 8649.
(62) Osakada, Y.; Kawai, K.; Fujitsuka, M.; Majima, T. Nuc. Acids Res. 2008, 36, 5562.
(63) Bhattacharya, P.K.; Cha, J.; Barton, J.K. Nuc. Acids Res. 2002, 30, 4740.
(64) Buzzeo, M.C.; Barton, J.K. Bioconjugate Chem. 2008, 19, 2110.
(65) Boal, A.K.; Barton, J.K. Bioconjugate Chem. 2005, 16, 312.
(66) Williams, T.T.; Dohno, C.; Stemp, E.D.A.; Barton, J.K. J. Am. Chem. Soc. 2004,
126, 8148.
(67) Furrer, E.; Giese, B. Helv. Chim. Acta 2003, 86, 3623.
(68) Genereux, J.C.; Augustyn, K.E.; Davis, M.L.; Shao, F.; Barton, J.K. J. Am. Chem.
Soc. 2008, 130, 15150.
(69) Joy, A.; Ghosh, A.K.; Schuster, G.B. J. Am. Chem. Soc. 2006, 128, 5346.
(70) Delaney, S.; Yoo, J.; Stemp, E.D.A.; Barton, J.K. Proc. Natl. Acad. Sci. U.S.A. 2004,
101, 10511.
(71) Nakatani, K.; Dohno, C.; Saito, I. J. Am. Chem. Soc. 2001, 123, 9681.
(72) O’Neill, M. A.; Dohno, C.; Barton, J. K. J. Am. Chem. Soc. 2004, 126, 1316.
(73) Dohno, C.; Ogawa, A.; Nakatani, K.; Saito, I. J. Am. Chem. Soc. 2003, 125, 10154.
(74) Elias, B.; Genereux, J.C.; Barton, J.K. Angew. Chem. Int. Ed. 2008, 47, 9067.
(75) Shao, F.; O’Neill, M. A.; Barton, J. K. Proc. Natl. Acad. Sci. U.S.A. 2004, 101,
17914.
(76) Shao, F.; Augustyn, K. E.; Barton, J. K. J. Am. Chem. Soc. 2005, 127, 17445.
74
(77) Liu, T.; Barton, J.K. J. Am. Chem. Soc. 2005, 127, 10160.
(78) Osakada, Y.; Kawai, K.; Fujitsuka, M.; Majima, T. Chem. Communications 2008,
23, 2656.
(79) Nakatani, K.; Dohno, C.; Saito, I. J. Am. Chem. Soc. 2000, 122, 5893.
(80) Kodera, H.; Osakada, Y.; Kawai, K.; Majima, T. Nat. Chem. 2009, 1, 156.
(81) Okamoto, A.; Nakatani, K.; Saito, I. J. Am. Chem. Soc. 2003, 125, 5066.
(82) Hunter, W.N.; Brown, T.; Kennard, O. Nuc. Acids. Res. 1987, 15, 6589.
(83) Hunter, W.N.; Brown, T.; Kneale, G.; Anand, N.N.; Rabinovich, D.; Kennard, O. J.
Biol. Chem., 1987, 262, 9962.
(84) Kumamoto, S.; Watanabe, M.; Kawakami, N.; Nakamura, M.; Yamanato, K.
Bioconjugate Chem. 2008, 19, 65.
(85) Takada, T.; Fujitsuka, M.; Majima, T. Proc. Natl. Acad. Sci. U.S.A. 2007, 104,
11179.
(86) Elias, B.; Shao, F.; Barton, J. K. J. Am. Chem. Soc. 2008, 130, 1152.
(87) Giese, B.; Wessely, S. Chem. Communications 2001, 20, 2108.
(88) Boal, A. K.; Yavin, E.; Lukianova, O. A.; O’Shea, V. L.; David, S. S.; Barton, J. K.
Biochemistry. 2005, 44, 8397.
(89) Gasper, S.M.; Schuster, G.B. J. Am. Chem. Soc. 1997, 119, 12762.
(90) Hickerson, R.P.; Prat, F.; Muller, J.G.; Foote, C.S.; Burrows, C.J. J. Am. Chem. Soc.
1999, 121, 9423.
(91) Drummond, T.G.; Hill, M.G.; Barton, J.K. Nat. Biotech. 2003, 21, 1192.
75
(92) Inouye, M.; Ikeda, R.; Takase, M.; Tsuri, T.; Chiba, J. Proc. Natl. Acad. Sci. U.S.A.
2005, 102, 11606.
(93) Boal, A.K.; Yavin, E.; Barton, J.K. J. Inorg. Biochem. 2007, 101, 1913.
(94) Gorodetsky, A.A.; Dietrich, L.E.P.; Lee, P.E.; Demple, B.; Newman, D.K.; Barton,
J.K. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 3684.
(95) Watanabe, S.; Kita, A.; Kobayashi, K.; Miki, K. Proc. Natl. Acad. Sci. U.S.A. 2008,
105, 4121.
(96) Gorodetsky, A.A.; Ebrahim, A.; Barton, J.K. J. Am. Chem. Soc. 2008, 130, 2924.
(97) Rajski, S. R.; Barton, J. K., Biochemistry 2001, 40, 5556.
(98) Nakatani, K.; Dohno, C.; Ogawa, A.; Saito, I. Chem. Biol. 2002, 9, 361.
(99) Kurbanyan, K.; Nguyen, K.L.; To, P.; Rivas, E.V.; Lueras, A.M.K.; Kosinski, C.;
Steryo, M.; González, A.; Mah, D.A.; Stemp, E.D.A. Biochemistry 2003, 42, 10269.
(100) Perrier S.; Hau, J.; Gasparutto, D.; Cadet, J.; Favier, A.; Ravanat, J.L. J. Am. Chem.
Soc. 2006, 128, 5703.
(101) Cadet, J.; Douki, T.; Ravanat, J.-L. Acc. Chem. Res. 2008, 41, 1075.
(102) Bjorklund, C.C.; Davis, W.B. Biochemistry 2007, 46, 10745.
(103) Nuñez, M.E.; Noyes, K.T.; Barton, J.K. Chem. Biol. 2002, 9, 403.
(104) Nuñez, M.E.; Holmquist, G.P.; Barton, J.K. Biochemistry 2001, 40, 12465.
(105) Merino, E.J.; Barton, J.K. Biochemistry 2008, 47, 1511.
(106) Nguyen, K.L.; Steryo, M.; Kurbanyan, K.; Nowitzki, K.M.; Butterfield, S.M.;
Ward, S.R.; Stemp, E.D.A. J. Am. Chem. Soc. 2000, 122, 3585.
76
(107) Johansen, M.E.; Muller, J.G.; Xu, X.Y.; Burrows, C.J. Biochemistry 2005, 44,
5660.
(108) Peng, X.; Pigli, Y.Z.; Rice, P.A.; Greenberg, M.M. J. Am. Chem. Soc. 2008, 130,
12890.
(109) Liu, C.-S. Schuster, G.B. J. Am. Chem. Soc. 2003, 125, 6098.
(110) Lewis, F.D.; Daublain, P.; Cohen, B.; Vura-Weiss, J.; Shafirovich, V.;
Wasielewski, M.R. J. Am. Chem. Soc. 2007, 129, 15130.
(111) O’Neill, M.A.; Barton, J.K. Proc. Natl. Acad. Sci U.S.A. 2002, 99, 16543.
(112) Shao, F.; Barton, J.K. J. Am. Chem. Soc. 2007, 129, 14733.
(113) Grozema, F.C.; Tonzani, S.; Berlin, Y.A.; Schatz, G.C.; Siebbeles, L.D.A.; Ratner,
M.A. J. Am. Chem. Soc. 2008, 130, 5157.
(114) Davis, W.B.; Ratner, M.A.; Wasielewski, M.R. J. Am. Chem. Soc. 2001, 123, 7877.
(115) Smalley, J.F.; Sachs, S.B.; Chidsey, C.E.D.; Dudek, S.P.; Sikes, H.D.; Creager,
S.E.; Yu, C.J.; Feldberg, S.W.; Newton, M.D. J. Am. Chem. Soc. 2004, 126, 14620.
(116) Newton, M.D.; Smalley, J.F. Phys. Chem. Chem. Phys. 2007, 9, 555.
(117) O’Neill, M. A.; Barton, J. K. J. Am. Chem. Soc. 2004, 126, 13234.
(118) Siriwong, K.; Voityuk, A.A.; Newton, M.D.; Rösch, N. J. Phys. Chem. B 2003,
107, 2595.
(119) Kocherzhenko, A.A., Patwardhan, S.; Grozema, F.C.; Anderson, H.L.; Siebbeles,
L.D.A. J. Am. Chem. Soc. 2009, 131, 5522.
(120) Wan, C.Z.; Fiebig, T.; Kelley, S.O.; Treadway, C.R.; Barton, J.K.; Zewail, A.H.
Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 6014.
77
(121) Kelley, S.O.; Holmlin, R.E.; Stemp, E.D.A.; Barton, J. K. J. Am. Chem. Soc. 1997,
199, 9861.
(122) Lewis, F.D.; Wu, T.; Xhang, Y.; Letsinger, R.L.; Greenfield, M.R.; Wasielewski,
M.R. Science 1997, 277, 673.
(123) Lewis, F.D.; Wu, T.; Liu, X.; Letsinger, R.L.; Greenfield, S.R.; Miller, S.E.;
Wasielewski, M.R. J. Am. Chem. Soc. 2000, 122, 2889.
(124) Lewis, F.D.; Zhu, H.; Daublain, P.; Fiebig, T.; Raytchev, M.; Wang, Q.;
Shafirovich, V. J. Am. Chem. Soc. 2006, 128, 791.
(125) Lewis, F.D.; Zhu, H.; Daublain, P.; Cohen, B.; Wasielewski, M.R. Angew. Chem.
Int. Ed. 2006, 45, 7982.
(126) Lewis, F.D.; Zhu, H.; Daublain, P.; Sigmund, K.; Fiebig, T.; Raytchev, M.; Wang,
Q.; Shafirovich, V. Photochem. Photobiol. Sciences 2008, 7, 534.
(127) Andreatta, D.; Pérez Lustres, J.L.; Kovalenko, S.A.; Ernsting, N.P.; Murphy, C.J.;
Coleman, R.S.; Berg, M.A. J. Am. Chem. Soc. 2005, 127, 7270.
(128) Dohno, C.; Stemp, E. D. A.; Barton, J. K. J. Am. Chem. Soc. 2003, 125, 9586.
(129) Armitage, B.; Yu, C.; Devadoss, C.; Schuster, G.B. J. Am. Chem. Soc. 1994, 116,
9847.
(130) Sanii, L.; Schuster, G.B. J. Am. Chem. Soc. 2000, 122, 11545.
(131) Vicic, D.A.; Odom, D.T.; Núñez, M.E.; Gianolio, D.A.; McLaughlin, L.W.;
Barton, J.K. J. Am. Chem. Soc. 2000, 122, 8603.
(132) Dotse, A.K.; Boone, E.K.; Schuster, G.B. J. Am. Chem. Soc. 2000, 122, 6825.
78
(133) Kawai, K.; Osakada, Y.; Takada, T.; Fujitsuka, M.; Majima, T. J. Am. Chem. Soc.
2004, 126, 12843.
(134) Kawai, K.; Osakada, Y.; Fujitsuka, M.; Majima, T. Chem. Eur. J. 2008, 14, 3721.
(135) Kawai, K.; Takada, T.; Nagai, T.; Cai, X.; Sugimoto, A.; Fujitsuka, M. ; Majima,
T. J. Am. Chem. Soc. 2003, 125, 16198.
(136) Kawai, K.; Osakada, Y.; Fujitsuka, M.; Majima, T. J. Phys. Chem. B 2008, 112,
2144.
(137) Arkin, M.R.; Stemp, E.D.A.; Holmlin, R.E.; Barton, J.K.; Hörmann, A.; Olson,
E.J.C.; Barbara, P.F. Science 1996, 273, 475.
(138) Wan, C.Z.; Fiebig, T.; Schiemann, O.; Barton, J.K., Zewail, A.H. Proc. Natl. Acad.
Sci. U.S.A. 2000, 97, 14052.
(139) Lewis, F.D.; Liu, J.; Zuo, X.; Hayes, R.T.; Wasielewski, M.R. J. Am. Chem. Soc.
2003, 125, 4850.
(140) Senthilkumar, K.; Grozema, F.C.; Guerra, C.F.; Bilkelhaupt, F.M.; Lewis, F.D.;
Berlin, Y.A.; Ratner, M.A.; Siebbeles, L.D.A. J. Am. Chem. Soc. 2005, 127, 14894.
(141) Santhosh, U.; Schuster, G.B. J. Am. Chem. Soc. 2002, 124, 10986.
(142) O’Neill, M.A.; Barton, J.K. In Topics in Current Chemistry; Schuster, G.B., Ed.;
Springer: Berlin, 2004; Vol. 237, p. 67.
(143) Boussicault, F.; Robert, M. Chem. Rev. 2008, 108, 2622.
(144) Seidel, C.A.M.; Shulz, A.; Sauer, M.H.M. J. Phys. Chem. 1996, 100, 5541.
(145) Caruso, T.; Carotenuto, M.; Vasca, E.; Peluso, A. J. Am. Chem. Soc. 2005, 127,
15040.
79
(146) Sheu, C.; Foote, C.S. J. Am. Chem. Soc. 1995, 117, 6439.
(147) Thorp, H.H. In Topics in Current Chemistry; Schuster, G.B., Ed.; Springer: Berlin,
2004; Vol. 237, p. 159.
(148) Johnston, D.H.; Glasgow, K.C.; Thorp, H.H. J. Am. Chem. Soc. 1995, 117, 8933.
(149) Shinde, S.S.; Maroz, A.; Hay, M.P.; Anderson, R.F. J. Am. Chem. Soc. 2009, 131,
5203.
(150) Sistare, M.F.; Codden, S.J.; Heimlich, G.; Thorp, H.H. J. Am. Chem. Soc. 2000,
122, 4742.
(151) Lewis F.D.; Liu, X.; Hayes, R.T.; Wasielewski, M.R. J. Am. Chem. Soc. 2000, 122,
12037.
(152) Kurnikov, I.V.; Tong, G.S.M.; Madrid, M.; Beratan, D.N. J. Phys. Chem. B 2002,
106, 7.
(153) Saito, I.; Takayama, M.; Sugiyama, H.; Nakatani, K. J. Am. Chem. Soc. 1995, 117,
6406.
(154) Sugiyama, H.; Saito, I. J. Am. Chem. Soc. 1996, 118, 7063.
(155) Adhikary, A.; Khanduri, D.; Sevilla, M.D. J. Am. Chem. Soc. 2009, 131, 8614.
(156) Hall, D.B.; Holmlin, R.E.; Barton, J.K. Nature 1996, 382, 731.
(157) Saito, I.; Nakamura, T.; Nakatani, K.; Yoshioka, Y.; Yamaguchi, K.; Sugiyama, H.
J. Am. Chem. Soc. 1998, 120, 12686.
(158) Voityuk, A.A. ; Jortner, J. ; Bixon, M. ; Rösch, N. Chem. Phys. Lett. 2000, 324,
430.
(159) Dandliker, P.J. ; Nuñez, M.E. ; Barton, J.K. Biochemistry 1998, 37, 6491.
80
(160) Yavin, E.; Stemp, E.D.A.; O’Shea, V.L.; David, S.S.; Barton, J.K. Proc. Natl.
Acad. Sci. U.S.A. 2006, 3610.
(161) Treadway, C.R.; Hill, M.G.; Barton, J.K. Chem. Phys. 2002, 281, 409.
(162) Anne, A.; Demaille, C. J. Am. Chem. Soc. 2008, 130, 9822.
(163) Wong, E.L.S.; Gooding, J.J. Anal. Chem. 2006 78, 2138.
(164) Gorodetsky, A.A.; Buzzeo, M.C.; Barton, J.K. Bioconjugate Chem. 2008, 19, 2285.
(165) Ikeda, R.; Akaishi, A.; Chiba, J.; Inouye, M. ChemBioChem 2007, 8, 2219.
(166) Hartwich, G.; Caruana, D.J.; de Lumley-Woodyear, T.; Wu, Y.; Campbell, C.N.;
Heller, A. J. Am. Chem. Soc. 1999, 121, 10803.
(167) Lewis, F.D.; Kalgutkar, R.S.; Wu, Y.; Liu, X.; Liu, J.; Hayes, R.T.; Miller, S.E.;
Wasielewski, M.R. J. Am. Chem. Soc. 2000, 122, 12346.
(168) LeBard, D.N.; Lilichenko, M.; Matyushov, D.V.; Berlin, Y.A.; Ratner, M.A. J.
Phys. Chem. B 2003, 107, 14509.
(169) Lewis, F.D.; Liu, J.; Weigel, W.; Rettig, W.; Kurnikov, I.V.; Beratan, D.N. Proc.
Natl. Acad. Sci. U.S.A. 2002, 99, 12536.
(170) Huynh, M.H.V.; Meyer, T.J. Chem. Rev. 2007, 107, 5004.
(171) Stubbe, J.; Nocera, D.G.; Yee, C.S.; Chang, M.C.Y. Chem. Rev. 2003, 103, 2167.
(172) Reece, S.Y.; Hodgkiss, J.M.; Stubbe, J.; Nocera, D.G. Phil. Trans. R. Soc. B. 2006,
361, 1351.
(173) Seyedsayamdost, M.R.; Yee, C.S.; Reece, S.Y.; Nocera, D.N.; Stubbe, J. J. Am.
Chem. Soc. 2006, 128, 1562.
(174) Fiebig, T.; Wan, C.; Zewail, A.H. ChemPhysChem 2002, 3, 781.
81
(175) Gervasio, F.L.; Boero, M.; Parrinello, M. Angew. Chem. Int. Ed. 2006, 45, 5606.
(176) Anderson, R.F.; Shinde, S.S.; Maroz, A. J. Am. Chem. Soc. 2006, 128, 15966.
(177) Kobayashi, K.; Yamagami, R.; Tagawa, S. J. Phys. Chem. B 2008, 112, 10752.
(178) Yamagami, R.; Kobayashi, K.; Tagawa, S. J. Am. Chem. Soc. 2008, 130, 14772.
(179) Huber, R.; Fiebig, T.; Wagenknecht, H.-A. Chem. Comm. 2003, 15, 1878.
(180) Stemp, E.D.A.; Arkin, M.; Barton, J.K. J. Am. Chem. Soc. 1997, 119, 2921.
(181) Schiemann, O.; Turro, N.J.; Barton, J.K. J. Phys. Chem. B 2000, 104, 7214.
(182) Kumar, A.; Sevilla, M.D. J. Phys. Chem. B 2009, 113, 11359.
(183) Shafirovich, V.; Dourandin, A.; Geacintov, N.E. J. Phys. Chem. B 2001, 105, 8431.
(184) Takada, T.; Kawai, K.; Fujitsuka, M.; Majima, T. Chem. Eur. J. 2005, 11, 3835.
(185) Sobolewski, A.L.; Domcke, W. Phys. Chem. Chem. Phys. 2004, 6, 2763.
(186) Sobolewski, A.L.; Domcke, W.; Hattig, C. Proc. Natl. Acad. Sci. U.S.A. 2005, 102,
17903.
(187) Abu-Riziq, A.; Grace, L., Nir, E.; Kabelac, M. ; Hobza, P. ; de Vries, M.S. Proc.
Natl. Acad. Sci. U.S.A. 2005, 102, 20.
(188) Schwalb, N.K.; Temps F. J. Am. Chem. Soc. 2007, 129, 9272.
(189) Crespo-Hernández, C.E.; de La Harpe, K.; Kohler, B. J. Am. Chem. Soc. 2008,
130, 10844.
(190) Brillouin, L. in Horizons in Biochemistry 1962 ed. M. Kasha and B. Pullman
Academic Press: New York City, 1962, 295.
(191) Sekiguchi, I.-S.; Sekiguchi, T. Phys. Rev. Lett. 2007, 99, 228102.
82
(192) Baba, Y.; Sekiguchi, T.; Shimoyama, I.; Hirao, N.; Nath, K.G. Phys. Rev. B 2006,
74, 205433.
(193) Zakjevskii, V.V.; King, S.J.; Dolgounitcheva, O.; Zakrzewski, V.G.; Ortiz, J.V. J.
Am. Chem. Soc. 2006, 128, 13350.
(194) Close, D.M.; Øhman, K.T. J. Phys. Chem. A 2008, 112, 11207.
(195) Gabelica, V.; Rosu, F.; Tabarin, T.; Kinet, C.; Rodolphe, A.; Broyer, M.; De Pauw,
E.; Dugourd, P. J. Am. Chem. Soc. 2007, 129, 4706.
(196) Hübsch, A.; Enders, R.G.; Cox, D.L.; Singh, R.R.P. Phys. Rev. Lett. 2005, 94,
178102.
(197) Haruna, K.-I.; Iida, H.; Tanabe, K.; Nishimoto, S.-I. Org. Biomolec. Chem. 2008, 6,
1613.
(198) Vladimirov, E.; Ivanova, A.; Rösch, N. J. Phys. Chem. B 2009, 113, 4425.
(199) Yonemoto, E.H.; Saupe, G.B.; Schmehl, R.H.; Hubig, S.M.; Riley, R.L.; Iverson,
B.L.; Mallouk, T.E. J. Am. Chem. Soc. 1994, 116, 4786.
(200) Berlin, Y.A.; Grozema, F.C.; Siebbeles, L.D.A.; Ratner, M.A. J. Phys. Chem. C
2008, 112, 10988.
(201) Page, C.C.; Moser, C.C.; Chen, X.; Dutton, P.L. Nature 1999, 402, 47.
(202) Gray, H.B.; Winkler, J.R. Q. Rev. Biophys. 2003, 36, 341.
(203) Balabin, I.A.; Beratan, D.N.; Skourtis, S.S. Phys. Rev. Lett. 2008, 101, 158102.
(204) Rösch, N.; Voityuk, A.A. In Topics in Current Chemistry; Schuster, G.B., Ed.;
Springer: Berlin, 2004; Vol. 237, p. 37.
(205) Sponer, J.; Riley, K.E.; Hobza, P. Phys. Chem. Chem. Phys. 2008, 10, 2595.
83
(206) Wang, X.F.; Chakraborty, T. Phys. Rev. Lett. 2006, 97, 106602.
(207) Giese, B.; Amaudrut; Köhler, A.-K.; Sporman, M.; Wessely, S. Nature 2001, 412,
318.
(208) Voityuk, A.A. J. Chem. Phys. 2008, 128, 115101.
(209) Migliore, A.; Corni, S.; Varsano, D.; Klein, M.L.; Di Felice, R. J. Phys. Chem. B
2009, 113, 9402.
(210) Troisi, A.; Orlandi, G. Chem. Phys. Lett. 2001, 344, 509.
(211) Troisi, A.; Orlandi, G. J. Phys. Chem. B 2002, 106, 2093.
(212) Mallajosyula, S.S.; Gupta, A.; Pati, S.K. J. Phys. Chem. A 2009, 113, 3955.
(213) Reha, D.; Barford, W.; Harris, S. Phys. Chem. Chem. Phys. 2008, 10, 5437.
(214) Kuba , T.; Elstner, M.; J. Phys. Chem. B 2008, 112, 8788.
(215) Bongiorno, A. J. Phys. Chem. B 2008, 112, 13945.
(216) Kuba , T.; Elstner, M.; J. Phys. Chem. B 2009, 113, 5653.
(217) Renger, T., Marcus, R. A. J. Phys. Chem. A 2003, 107, 8404.
(218) Shih, C.; Museth, A.K.; Abrahamsson, M.; Blanco-Rodriguez, A.M.; Di Bilio, A.J.;
Sudhamsu, J.; Crane, B.R.; Ronayne, K.L.; Towrie, M.; Vicek, A.; Richards, J.H.;
Winkler, J.R.; Gray, H.B. Science 2008, 320, 1760.
(219) Jorner, J.; Bixon, M.; Langenbacher, T.; Michel-Beyerle, M.E. Proc. Natl. Acad.
Sci U.S.A. 1998, 95, 12759.
(220) Xu, B.; Tao, N. Science 2003, 301, 1221.
(221) Hihath, J.; Chen, F.; Zhang, P.; Tao, N. J. Phys. Condens. Matter 2007, 19,
215202.
84
(222) Mallajosyula, S.S.; Lin, J.C.; Cox, D.L.; Pati, S.K.; Singh, R.R.P. Phys. Rev. Lett.
2008, 101, 176805.
(223) Hihath, J.; Xu, B.; Zhang, P.; Tao, N. Proc. Natl. Acad. Sci. U.S.A. 2005, 102,
16979.
(224) Dulic, D., Tuukkanen, S., Chung, C.-L.; Isambert, A.; Lavie, P.; Filoramo, A.
Nanotech. 2009, 20, 115502.
(225) Barton, J.K.; Nuckols, C. Unpublished results.
(226) Takada, T.; Kawai, K.; Cai, X.; Sugimoto, A.; Fujitsuka, M.; Majima, T. J. Am.
Chem. Soc. 2004, 126, 1125.
(227) Takada, T.; Kawai, K.; Fujitsuka, M.; Majima, T.; J. Am. Chem. Soc. 2006, 128,
11012.
(228) Jortner, J.; Bixon, M.; Voityuk, A.A.; Rösch, N. J. Phys. Chem. A 2002, 106, 7599.
(229) Sartor, V.; Boone, E.; Schuster, G.B. J. Phys. Chem. B 2001, 105, 11057.
(230) Yoo, J.; Delaney, S.; Stemp, E.D.A.; Barton, J.K. J. Am. Chem. Soc. 2003, 125,
6640.
(231) Kendrick, T.; Giese, B. Chem. Comm. 2002, 18, 2016.
(232) Giese, B.; Wessely, S. Angew. Chem. Int. Ed. 2000, 39, 3490.
(233) Bixon, M.; Jortner, J. J. Am. Chem. Soc. 2001, 123, 12556.
(234) Bixon, M.; Jortner, J. Chem. Phys. 2002, 271, 393.
(235) Bixon, M.; Jortner, J. Chem. Phys. 2006, 326, 252.
(236) Liu, C.-S.; Hernandez, R.; Schuster, G.B. J. Am. Chem. Soc. 2004, 126, 2877.
85
(237) Goldsmith, R.H.; DeLeon, O.; Wilson, T.M.; Finkelstein-Shapiro, D.; Ratner,
M.A.; Wasielewski, M.R. J. Phys. Chem. A 2008, 112, 4410.
(238) Yu, Z.G.; Song, X. Phys. Rev. Lett. 2001, 86, 6018.
(239) Berlin, Y.A.; Burin, A.L.; Ratner, M.A. J. Am. Chem. Soc. 2001, 123, 260.
(240) Kelley, S. O.; Jackson, N. M.; Hill, M. G.; Barton, J. K. Angew. Chem., Int. Ed.
1999, 38, 941.
(241) Hartwich, G.; Caruana, D.J.; de Lumley-Woodyear, T.; Wu, Y.; Campbell, C.N.;
Heller, A. J. Am. Chem. Soc. 1999, 121, 10803.
(242) Traub, M.C.; Brunschwig, B.S.; Lewis, N.S. J. Phys. Chem. B 2007, 111, 6676.
(243) Adhikary, A.; Kumar, A.; Khanduri, D.; Sevilla, M.D. J. Am. Chem. Soc. 2008,
130, 10282.
(244) Jakobsson, M.; Stafström, S. J. Chem. Phys. 2008, 129, 125102.
(245) Schuster, G.B. Acc. Chem. Res. 2000, 33, 253.
(246) Conwell, E. M.; Rakhmanova, S. V. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 4556.
(247) Kuznetsov, A.M. Charge Transfer in Physics, Chemistry and Biology: Physical
Mechanisms of Elementary Processes and an Introduction to the Theory Gordon and
Breach Science Publishers: Amsterdam, 1995, 2.
(248) Maniadis, P.; Kalosakas, G; Rasmussen, K.Ø.; Bishop, A.R. Phys. Rev. B. 2003,
68, 174304.
(249) Komineas, S.; Kalosakas, G.; Bishop, A.R. Phys. Rev. E 2002, 65, 061905.
(250) Chang, C.-M.; Castro Neto, A.H.; Bishop, A.R. Chem. Phys. 2004, 303, 189.
(251) Conwell, E.M. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 8795.
86
(252) Conwell, E. M.; Bloch, S. M. J. Phys Chem. B 2006, 110, 5801.
(253) Conwell, E.M.; Park, J.-H.; Choi, H.-Y. J. Phys. Chem. B 2005, 109, 9760.
(254) Conwell, E.M.; Bloch, S.M.; McLaughlin, P.M.; Basko, D.M. J. Am. Chem. Soc.
2007, 129, 9175.
(255) Conwell, E.M.; Basko, D.M. J. Phys. Chem. B 2006, 110, 23603.
(256) Middleton, C.T.; de La Harpe, K.; Su, C. ; Law, Y.K.; Crespo-Hernández, C.E.;
Kohler, B. Ann. Rev. Phys. Chem. 2009, 60, 217.
(257) Schwalb, N.K.; Temps, F. Science 2008, 322, 243.
(258) Crespo-Hernández, C.E.; Cohen, B.; Kohler, B. Nature 2005, 436, 1141.
(259) Buchvarov, I.; Wang, Q.; Raytchev, M.; Trifonov, I.; Fiebig, T. Proc. Natl. Acad.
Sci. U.S.A. 2007, 104, 4794.
(260) Kadhane, U.; Holm, A.I.S.; Hoffmann, S.V.; Nielsen, S. Phys. Rev. E 2008, 77,
021901.
(261) Takaya, T,; Su, C.; de La Harpe, K.; Crespo-Hernández, C.E.; Kohler, B. Proc.
Natl. Acad. Sci. U.S.A. 2008, 105, 10285.
(262) Lange, A.W.; Herbert, J.M. J. Am. Chem. Soc. 2009, 131, 3913.
(263) Hatcher, E.; Balaeff, A.; Keinan, S.; Venkatramani, R.; Beratan, D.N. J. Am. Chem.
Soc. 2008, 130, 11752.
(264) Uskov, D.B.; Burin, A.L. Phys. Rev. B 2008, 78, 073106.
(265) Voityuk, A.A. J. Chem. Phys. 2005, 122, 204904.
(266) Sen, S.; Andreatta, D.; Ponomarev, S.Y.; Beveridge, D.L.; Berg, M.A. J. Am.
Chem. Soc. 2009, 131, 1724,
87
(267) Voityuk, A.A. J. Chem. Phys. 2008, 128, 045104.
(268) Friedman, K.A.; Heller, A. J. Phys. Chem. B 2001, 105, 11859.
(269) Heller, A. Farad. Disc. 2000, 116, 1.
(270) Merino, E.J.; Barton, J.K. Biochemistry 2007, 46, 2805.
(271) Merino, E.J.; Barton, J.K. Biochemistry 2009, 48, 660.
(272) Steenken, S.; Jovanovic, S.V. J. Am. Chem. Soc. 1997, 119, 617.
(273) Crespo-Hernández, C.E.; Close, D.M.; Gorb, L.; Leszczynski, J. J. Phys. Chem. B
2007, 111, 5386.